Skip to main content
NCBI home page
Acta Crystallographica Section D: Structural Biology logoLink to Acta Crystallographica Section D: Structural Biology
. 2019 Jul 30;75(Pt 8):733–742. doi: 10.1107/S2059798319009574

Biochemical and structural explorations of α-hydroxyacid oxidases reveal a four-electron oxidative decarboxylation reaction

Hsien-Wei Yeh a,‡, Kuan-Hung Lin a,b,‡, Syue-Yi Lyu a, Yi-Shan Li a, Chun-Man Huang a, Yung-Lin Wang a, Hao-Wei Shih a, Ning-Shian Hsu a, Chang-Jer Wu c, Tsung-Lin Li a,d,*
PMCID: PMC6677016  PMID: 31373572

Structural and enzymological explorations of α-hydroxyacid oxidases uncover new flavin mononucleotide-mediated reactions and intermediates.

Keywords: mandelate oxidase, flavin mononucleotide, oxidative decarboxylation, hydride transfer, α-hydroxyacids

Abstract

p-Hydroxymandelate oxidase (Hmo) is a flavin mononucleotide (FMN)-dependent enzyme that oxidizes mandelate to benzoylformate. How the FMN-dependent oxidation is executed by Hmo remains unclear at the molecular level. A continuum of snapshots from crystal structures of Hmo and its mutants in complex with physiological/nonphysiological substrates, products and inhibitors provides a rationale for its substrate enantioselectivity/promiscuity, its active-site geometry/reactivity and its direct hydride-transfer mechanism. A single mutant, Y128F, that extends the two-electron oxidation reaction to a four-electron oxidative decarboxylation reaction was unexpectedly observed. Biochemical and structural approaches, including biochemistry, kinetics, stable isotope labeling and X-ray crystallo­graphy, were exploited to reach these conclusions and provide additional insights.

1. Introduction  

l-p-Hydroxyphenylglycine (l-p-HPG), a nonproteinogenic α-amino acid, exists in many clinically important glycopeptide natural products, for example vancomycin (van Wageningen et al., 1998 ▸), teicoplanin (Li et al., 2004 ▸) and ramoplanin (McCafferty et al., 2002 ▸; Walker et al., 2005 ▸). The peptide cores of these glycopeptides recruit two, three or six l-p-HPG units, respectively, as the site for glycosylation in addition to their structural roles (Supplementary Fig. S1). The biosynthesis of l-p-HPG requires three enzymes, (S)-p-hydroxymandelate synthase (HmaS), (S)-p-hydroxymandelate oxidase (Hmo) and l-p-hydroxyphenylglycine transaminase (HpgT): HmaS transforms phenylpyruvate to (S)-mandelate, Hmo catalyzes the conversion of (S)-mandelate to benzoylformate and HpgT then transaminates benzoylformate to l-p-HPG [Fig. 1 ▸(a)] (Al Toma et al., 2015 ▸; Hubbard et al., 2000 ▸; Li et al., 2001 ▸). In the mandelate metabolism, (R)-mandelate is converted to (S)-mandelate by mandelate racemase, and (S)-mandelate is then oxidized to benzoylformate by (S)-mandelate dehydrogenase (MDH/MdlB). Benzoylformate subsequently undergoes decarboxylation to benzaldehyde by a thiamine-pyrophos­phate (ThDP)-dependent decarboxylase. Benzaldehyde is further oxidized by dehydrogenases to benzoate [Fig. 1 ▸(b)] (Petsko et al., 1993 ▸; Tsou et al., 1990 ▸).

Figure 1.

Figure 1

Open in a new tab

Pathways and structures. (a) Anabolic pathway of l-p-hydroxyphenylglycine. (b) Metabolic pathway of (R)-mandelate; MdlB is also referred to as S-­mandelate dehydrogenase (MDH). (c) Chemical structure of oxidized flavin mononucleotide (FMNox). (d) The direct conversion of mandelate to benzoate by the Y128F mutant via a four-electron oxidative decarboxylation reaction. The numbering system is shown in red for FMN and phenylpyruvate; the si-face and re-face prochirality relative to the N5 atom of the isoalloxazine ring is indicated in blue and green, respectively.

Hmo shares high sequence homology with flavin mononucleotide (FMN)-dependent oxidoreductases (Supplementary Fig. S2) such as MDH (Lehoux & Mitra, 1999 ▸), l-lactate oxidase (LO; Lockridge et al., 1972 ▸), flavocytochrome b 2 (FCB2; Xia & Mathews, 1990 ▸), glycolate oxidase (GO; Lindqvist, 1989 ▸), l-lactate monooxygenase (LMO; Giegel et al., 1990 ▸; Maeda-Yorita et al., 1995 ▸), long-chain l-α-hydroxy­acid oxidase (LCHAO; Cunane et al., 2005 ▸), human glycolate oxidase (hGOX; Murray et al., 2008 ▸) and human l-α-hydroxyacid oxidase 2 (HAOX2; Jones et al., 2000 ▸). With the exception of LMO, all of these oxidases execute two-electron oxidation of α-hydroxyacids to the corresponding α-ketoacids following the classic ping-pong reaction mechanism. In contrast to β-ketoacids, α-ketoacids are devoid of a suitably placed electron sink to stabilize the negative charge developed upon decarboxylation. For the decarboxylation of α-ketoacids, a common solution in nature is to utilize the cofactor ThDP or pyridoxal phosphate (PLP), as exemplified in the central metabolism (Perham et al., 2002 ▸). To the best of our knowledge, LMO is the only ThDP/PLP/NADPH-independent enzyme that performs oxidative decarboxylation of l-lactate en route from pyruvate to acetate at the expense of one molecule of O2, with the concomitant production of CO2 and H2O, and thus is referred to as a monooxygenase (Ghisla & Massey, 1989 ▸). When lactate is oxidized to pyruvate, the oxidized FMN [FMNox; Fig. 1 ▸(c)] is concertedly reduced to FMNred upon receiving a pair of electrons: the reductive half-reaction. FMNred is instantly oxidized to FMNox by O2 to form hydrogen peroxide (H2O2): the oxidative half-reaction (Fagan & Palfey, 2010 ▸; Romero et al., 2018 ▸). Given that the dissociation of pyruvate from LMO is a slow step, the H2O2 generated at the active site acts on pyruvate to form acetate via oxidative decarboxylation (Giegel et al., 1990 ▸; Lopalco et al., 2016 ▸). Aside from this non-ping-pong kinetic description, how H2O2 mediates the oxidative decarboxylation of α-ketoacids remains elusive (Choong & Massey, 1980 ▸; Ghisla & Massey, 1977 ▸; Lockridge et al., 1972 ▸; Walsh et al., 1973 ▸). In this study, we explored Hmo and its mutants to gain insights into the reaction center [Fig. 1 ▸(d); see below]. A biochemical/structural biology approach renders various liganded structures of the proteins, which in combination deconvolve reactive/unstable Hmo and its mutant-mediated reactions at the molecular level.

2. Materials and methods  

2.1. Protein expression and purification  

The ligated plasmids of native hmo and its mutants were transformed into Escherichia coli BL21(DE3) cells and cultured in 1 l LB medium containing 50 mg l−1 kanamycin at 37°C until the A 600 reached 0.6. For protein expression and purification, 200 µl of 1.0 M isopropyl β-d-thiogalacto­pyranoside (IPTG) was added to the cultured E. coli medium for a further 12 h at 16°C. The cells were harvested by centrifugation, resuspended in 40 ml binding buffer (20 mM HEPES pH 7.5, 500 mM NaCl, 10 mM imidazole, 10% glycerol) and ruptured by sonication. Cell debris was spun down by centrifugation (20 000 rev min−1, 30 min, 4°C) and the supernatants were applied onto an Ni2+–NTA resin column for further purification. The column was washed twice with buffers at different concentrations (first wash, 20 mM HEPES pH 8.0, 500 mM NaCl, 40 mM imidazole; second wash, 20 mM HEPES pH 8.0, 500 mM NaCl, 80 mM imidazole) before eluting the target protein with 15 ml elution buffer (20 mM HEPES pH 8, 500 mM NaCl, 250 mM imidazole). All proteins were further purified by size-exclusion chromatography on an ÄKTA FPLC system equipped with a HiLoad 16/60 Superdex 200 column under an isocratic condition (20 mM HEPES pH 8.0, 100 mM NaCl). Protein purity was examined by SDS–PAGE. Protein concentrations were estimated by the Bradford assay using BSA as a standard.

2.2. Crystallization and data collection  

Hmo and its Y128F mutant were crystallized using the hanging-drop vapor-diffusion method at 20°C. Each protein was concentrated to 7 mg ml−1 in 50 mM HEPES pH 8.0 buffer solution and was then mixed with the same volume of reservoir solution (35% Tascimate, 0.1 M bis-Tris propane pH 7.0). Crystals generally formed after a five-day incubation. Before X-ray diffraction data collection, the crystals were transferred to the same mother liquor containing 20% glycerol as a cryoprotectant. For Hmo or its Y128F mutant in complex with 4-hydroxy-(S)-mandelate (4HSMA), (S)-mandelate (SMA), benzoylformate (BF), benzoic acid (BA), benzaldehyde (BZ), (S)-propionic acid (SPPA), l-4-hydroxyphenylglycine (LPG) and phenylpyruvate, X-ray data sets were recorded after soaking protein crystals with 10–30 mM of the given substrate for 10 min to 24 h in the reservoir solution. All diffraction data were collected using ADSC Quantum 315 or MX300-HE CCD detectors at an operating temperature of 100 K on beamlines 13B1, 13C1, 15A1 and 05A at the National Synchrotron Radiation Research Center (NSRRC), Taiwan or on beamline 44XU at SPring-8, Japan. Data were indexed, and scaled with the HKL-2000 package (Otwinowski & Minor, 1997 ▸). The Y128F mutant crystals belonged to space group I422, with unit-cell parameters a = 137.565, b = 137.565, c = 112.087 Å.

2.3. Structure determination and refinement  

The crystal phase information for Hmo was obtained by molecular replacement using Phaser-MR from the CCP4 suite (McCoy et al., 2007 ▸; Winn et al., 2011 ▸). Hydroxyacid oxidase (PDB entry 3sgz; Chen et al., 2012 ▸) was used as the search model to solve the initial phase. Polypeptide models were built and refined with REFMAC (Murshudov et al., 2011 ▸), Coot (Emsley et al., 2010 ▸) and PHENIX (Afonine et al., 2012 ▸). Detailed refinement statistics are presented in Table 1 ▸. Protein structures and electron-density maps were presented using PyMOL (http://www.pymol.org).

Table 1. Data-collection and refinement statistics for Hmo and its Y128F mutant.

Values in parentheses are for the highest resolution shell. 4HSMA, 4-hydroxy-(S)-mandelate; SMA, (S)-mandelate; BF, benzoylformate; BA, benzoate; SPPA, 2-­phenylpropanoate; BZ, benzaldehyde; PA, 2-phenylacetate; PLA, 2-hydroxy-3-phenylpropanoate; LLAC, 2-hydroxypropanoate; RMA, (R)-mandelate; PPY, phenylpyruvic acid.

  Hmo Hmo–4HSMA Hmo–SMA Hmo–BF Hmo–BA Hmo–SPPA Y128F Y128F–BZ Y128F–LLAC
PDB code 5zzp 5zzq 5zzr 6a08 5zzs 6a00 6a13 6a0o 5zzy
Data collection
 Wavelength (Å) 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0
 Space group I422 I422 I422 I422 I422 I422 I422 I422 I422
  a, b, c (Å) 137.9, 137.9, 112.3 137.8, 137.8, 111.3 137.7, 137.7, 111.6 137.9, 137.9, 111.8 137.9, 137.9, 111.8 138.2, 138.2, 111.7 137.5, 137.5, 112.0 137.7, 137.7, 112.0 137.8, 137.8, 111.9
 α, β, γ (°) 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90
 Resolution range (Å) 30–1.39 (1.44–1.39) 30–1.32 (1.37–1.32) 30–1.31 (1.36–1.31) 30–1.55 (1.61–1.55) 30–1.40 (1.45–1.40) 30–1.60 (1.66–1.60) 30–1.70 (1.76–1.70) 30–1.65 (1.71–1.65) 30–1.50 (1.55–1.50)
  R merge † (%) 3.7 (73.0) 3.8 (80.1) 3.7 (67.0) 4.0 (79.0) 3.9 (67.1) 4.8 (66.2) 3.5 (69.2) 3.9 (62.1) 5.4 (82.1)
 〈I/σ(I)〉 36.1 (2.4) 43.5 (2.5) 34.5 (2.3) 43.1 (2.9) 32.3 (2.9) 28.5 (2.1) 41.3 (3.1) 31.0 (2.2) 31.2 (2.0)
 Completeness (%) 99.9 (100.0) 100.0 (100.0) 99.0 (100.0) 100.0 (100.0) 99.0 (99.7) 99.5 (97.9) 100.0 (100.0) 100.0 (100.0) 99.9 (99.1)
 Multiplicity 9.4 (9.4) 12.2 (12.0) 9.8 (9.5) 12.8 (11.2) 9.7 (9.5) 8.8 (8.0) 11.2 (11.1) 9.5 (8.7) 11.6 (8.3)
Refinement
 Resolution range (Å) 30–1.39 (1.44–1.39) 30–1.32 (1.37–1.32) 30–1.31 (1.36–1.31) 30–1.55 (1.61–1.55) 30–1.40 (1.45–1.40) 30–1.60 (1.66–1.60) 30–1.70 (1.76–1.70) 30–1.65 (1.71–1.65) 30–1.50 (1.55–1.50)
  R work ‡ (%) 17.0 (28.0) 17.5 (23.7) 16.6 (22.9) 16.9 (22.0) 16.4 (23.1) 17.4 (26.8) 17.8 (24.2) 15.8 (20.6) 15.8 (25.6)
  R free § (%) 18.3 (30.0) 18.8 (25.3) 17.7 (22.3) 17.9 (20.1) 18.4 (25.0) 19.0 (30.8) 20.0 (26.2) 18.2 (22.1) 16.8 (25.8)
 R.m.s. deviations
  Bond lengths (Å) 0.010 0.010 0.009 0.008 0.015 0.010 0.018 0.010 0.023
  Bond angles (°) 1.41 1.42 1.41 1.26 1.37 1.223 1.490 1.446 2.119
 No. of reflections 100548 112024 112497 75720 102688 69702 58599 54457 78351
 No. of atoms
  Protein 2785 2817 2622 2684 2656 2651 2510 2659 2551
  Ligand/ion 31 43 53 65 58 53 31 58 37
  Water 403 321 402 329 368 307 316 364 341
  B factors (Å2)
  Protein 18.2 19.5 19.7 19.1 18.2 19.8 18.6 17.7 18.4
  Ligand/ion 10.7 15.6 19.0 20.6 18.9 26.9 10.8 19.4 14.6
  Water 32.4 30.1 33.9 31.7 32.8 31.6 32.1 32.6 32.2
Open in a new tab
  Y128F–RMA Y128F–4HSMA Y128F–SMA Y128F–BF Y128F–BA Y128F–PPY Y128F–PA Y128F–PLA Y128F–SPPA
PDB code 5zzx 6a1a 6a0v 6a19 6a0y 6a11 6a0m 6a0g 6a0d
Data collection
 Wavelength (Å) 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0
 Space group I422 I422 I422 I422 I422 I422 I422 I422 I422
  a, b, c (Å) 138.1, 138.1, 112.4 137.8, 137.8, 111.5 137.8, 137.8, 111.8 137.9, 137.9, 112.3 137.7, 137.7, 112.0 138.5, 138.5, 111.5 138.2, 138.2, 111.2 138.3, 138.3, 111.4 137.9, 137.9, 111.3
 α, β, γ (°) 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90 90, 90, 90
 Resolution range (Å) 30–1.50 (1.55–1.50) 30–1.35 (1.40–1.35) 30–1.39 (1.44–1.39) 30–1.55 (1.61–1.55) 30–1.65 (1.71–1.65) 30–1.45 (1.50–1.45) 30–1.75 (1.81–1.75) 30–1.80 (1.86–1.80) 30–1.65 (1.71–1.65)
  R merge † (%) 6.3 (78.0) 5.1 (74.6) 4.3 (73.1) 3.6 (75.0) 3.9 (62.1) 4.1 (79.0) 3.7 (79.0) 4.8 (81.0) 3.5 (71.0)
 〈I/σ(I)〉 27.5 (2.0) 36.09 (2.7) 41.0 (2.9) 32.9 (2.7) 31.0 (2.2) 42.0 (2.7) 35.8 (2.0) 34.5 (2.7) 35.9 (2.4)
 Completeness (%) 99.8 (100.0) 100.0 (100.0) 100.0 (100.0) 99.9 (100.0) 100.0 (100.0) 99.9 (100.0) 99.9 (100.0) 99.9 (100.0) 99.9 (100.0)
 Multiplicity 9.6 (9.3) 12.2 (12.0) 12.2 (12.0) 9.6 (9.2) 9.5 (8.7) 11.8 (11.9) 9.3 (7.8) 11.1 (10.7) 9.7 (9.2)
Refinement
 Resolution range (Å) 30–1.50 (1.55–1.50) 30–1.35 (1.40–1.35) 30–1.39 (1.44–1.39) 30–1.55 (1.61–1.55) 30–1.65 (1.71–1.65) 30–1.45 (1.50–1.45) 30–1.75 (1.81–1.75) 30–1.80 (1.86–1.80) 30–1.65 (1.71–1.65)
  R work ‡ (%) 17.5 (30.7) 16.7 (23.5) 16.7 (22.9) 15.7 (24.4) 15.8 (20.6) 16.9 (23.9) 16.4 (24.1) 16.0 (22.9) 11.7 (23.3)
  R free § (%) 19.9 (32.7) 18.0 (24.6) 18.4 (23.1) 17.3 (26.6) 18.2 (22.1) 19.1 (24.4) 18.5 (26.5) 18.2 (25.6) 19.1 (25.2)
 R.m.s. deviations
  Bond lengths (Å) 0.023 0.008 0.008 0.009 0.010 0.016 0.018 0.021 0.022
  Bond angles (°) 2.121 1.3811 1.3361 1.460 1.446 1.433 1.834 2.044 2.080
 No. of reflections 80736 106077 98014 74156 54457 91944 49594 46666 59471
 No. of atoms
  Protein 2547 2640 2636 2703 2659 2677 2555 2545 2530
  Ligand/ion 42 55 53 53 58 43 41 55 53
  Water 369 401 400 400 364 357 279 288 304
  B factors (Å2)
  Protein 17.7 16.8 16.9 21.9 17.7 20.0 25.8 25.3 20.2
  Ligand/ion 25.9 18.1 17.8 29.4 19.4 20.1 26.2 23.2 27.5
  Water 33.1 33.6 32.2 37.3 32.6 34.9 37.0 36.7 33.6
Open in a new tab
†

R merge = Inline graphic Inline graphic, where Ii(hkl) is the average intensity value of equivalent reflections.

‡

R work = Inline graphic Inline graphic.

§

R free was calculated from 5% of data that were randomly excluded from refinement.

2.4. Site-directed mutagenesis  

The hmo (orf22) gene was amplified from Amycolatopsis orientalis genomic DNA by PCR amplification and was then subcloned into the expression vector pET-28a(+) to produce an N-terminally His6-tagged protein. The expression plasmid pET-28a and PCR products were digested with the restriction enzymes NdeI and XhoI, respectively, at 37°C for 5 h. DNA ligation was subsequently performed according to the manufacturer’s instructions (Novagen). The inserted gene of the clones was checked by restriction-enzyme analysis and justified by agarose electrophoresis and DNA sequencing. The site-directed point mutants Y128A, Y128C, Y128F, R163L, H252A and R255A were generated using QuikChange (Stratagene), in which the wild-type hmo gene was used as the template for single mutants. All mutants were confirmed by DNA sequencing. Mutant proteins were purified using the same protocol as used for recombinant wild-type (WT) Hmo.

2.5. Enzymatic activity assay  

Commercial α-hydroxyacids/α-ketoacids, including (S)-p-hydroxymandelate, (S)-mandelate, p-chloromandelic acid, p-bromomandelic acid, 2-chloromandelic acid, 4-(trifluoro­methyl)mandelate and benzylformate, were purchased from Sigma–Aldrich. Other unusual substrates, including α-hydroxy­amides/α-ketoamides, were chemically prepared following modified synthetic protocols (see below). A typical enzymatic reaction was carried out in HEPES buffer (20 mM HEPES, 100 mM NaCl pH 7.5) using the substrates mentioned above and protein (native Hmo or mutants) at 25°C for 4 h. After quenching with 6 N HCl, the filtered reaction solutions were subjected to a reverse-phase C18 column (4.6 × 250 mm, 5 µm, C18 Prodigy, Phenomenex) mounted on an Agilent 1260 Infinity Quaternary LC module in connection with ESI/MS (Thermo-Finnigan LTQ-XL), a Waters Alliance 2695 HPLC module with a Xevo TQ-S micro triple quadrupole mass spectrometer, or GC-MS (Thermo-Finnigan Polaris Q). The LC analytical mobile phase was programmed as water with 1% formic acid as solvent A and acetonitrile with 1% formic acid as solvent B, with a linear gradient from 2% to 40% solvent B in solvent A at a flow rate of 1.0 ml min−1 over 25 min followed by 98% solvent B for another 8 min. The analytes were monitored at a UV wavelength at 254 nm using both positive and negative modes for mass detection. LC-MS data were recorded and analyzed using the MassLynx or Xcalibur software.

2.6. Measurement of the production of hydrogen peroxide  

Hydrogen peroxide production by Hmo and its mutants was measured using the Fluorescent Hydrogen Peroxide/Peroxidase Detection Kit (Fluoro H2O2). We followed the general protocol provided by Cell Technology and detected the fluorescence to quantify the production of hydrogen peroxide (excitation at 579 nm and emission at 600 nm).

2.7. Hmo and Y128F mutant reactions in the presence of 18O2 and H2 18O  

For the 18O2 experiments, the assay solution was made in a final volume of 200 µl that contained 20 mM HEPES, 100 mM NaCl, 2 mM (S)-mandelic acid and 0.4 mg Hmo Y128F mutant. The sample was subjected to vacuum and an N2 purge to remove O2. The sample was then refilled with 18O2. After 2 h of stirring at 25°C, the reaction mixture was denatured with 6 N HCl, centrifuged and analyzed by HPLC and GC-MS. For the H2 18O experiment, the Hmo Y128F mutant assays were carried out with the same reactants in a final buffer solution of 90% H2 18O. The enzymatic reactions were incubated at room temperature for 2 h and denatured with 6 N HCl. Both HPLC and GC-MS were used to detect the targets.

3. Results and discussion  

3.1. Reactivity of Hmo  

We first set out to examine Hmo from A. orientalis. Biochemical analysis showed that Hmo efficiently transforms (S)-mandelate to benzoylformate (Hubbard et al., 2000 ▸; Li et al., 2001 ▸), while a small peak that emerged at 17.5 min on LC traces in a dose-dependent manner drew our attention [Fig. 2 ▸(a)]. This peak was determined to be benzoate by NMR and MS (Supplementary Fig. S3). The enantiopreference for substrates is for (S)-conformers, as (R)-mandelate was left unchanged (Supplementary Fig. S4). The reactivity of Hmo towards the substrates (S)-mandelate, l-p-HPG and (S)-2-methylphenylacetate showed that the first two are oxidized to benzoylformate with benzoate as a minor product, but the third cannot be oxidized. The utilization of l-p-HPG shows that Hmo can act as an amine oxidase. The para and ortho substituents on the phenyl ring of (S)-mandelate have little influence on the activity of the enzyme, as 4-chloro-, 2-chloro-, 4-bromo- and 4-trifluoromethyl-(S)-mandelate can all be oxidized by Hmo to the corresponding α-ketoacids with benzoate as a minor product (Supplementary Fig. S4; Supplementary Table S2). The α-hydroxyacid moiety is not limited to two carbons: (S)-3-phenyllactate can be oxidized to phenylpyruvate with phenylacetate as a minor product (Supplementary Fig. S4). Although minor decarboxylated acid products can be formed, our analysis agreed that Hmo is an oxidase [k cat = 5.17 s−1, K m = 0.16 mM for the conversion of (S)-mandelate to benzoylformate].

Figure 2.

Figure 2

Open in a new tab

LC, UV and MS spectra for reactions catalyzed by Hmo and its mutants. (a) LC traces of enzymatic reactions with (S)-mandelate (1) as the substrate catalyzed by Hmo and its mutants: a, WT Hmo; b, Y128F mutant; c, R163L mutant; d, H252F mutant; e, R255A mutant; f, (S)-mandelate standard (1); g, benzoylformate standard (2). (b) Enzymatic reactions catalyzed by the Y128F mutant, where benzoate (3) is the major kinetic product over 420 min. (c) Mass spectra of methyl benzoate analyzed by GC-MS for enzymatic reactions conducted in the presence of H2 18O (left) and 18O2 (right). (d) Formation of hydrogen peroxide (H2O2) in enzymatic reactions catalyzed by WT Hmo and the Y128F mutant in the presence of (S)-mandelate (SMA) or 4-­hydroxy-(S)-mandelate (4HSMA). (e) LC traces of enzymatic reactions catalyzed by the R163L mutant with α-(S)-mandelic amide as the substrate (i), in which the product is 2-oxo-2-phenylacetamide showing the same retention as the synthetic reference (ii).

With respect to the mechanism of FMN-dependent oxidases, two mechanisms, a direct hydride-transfer mechanism (HT) and a carbanion mechanism (CA), have generally been referred to (Dijkman et al., 2013 ▸; Walsh & Wencewicz, 2013 ▸). For HT, deprotonation of the α-OH by an active-site base yields an oxyanion species, which upon collapse transfers an α-hydride to FMNox. For CA, the abstraction of a relatively acidic α-proton by an active-site base results in a dianionic/enolate intermediate, and electron transfer to this intermediate or covalent association with FMNox takes place. While the majority of relevant biochemical and kinetic studies support HT, unequivocal structural evidence remains lacking (Cao et al., 2014 ▸; Dijkman et al., 2013 ▸; Gillet et al., 2016 ▸; Lederer et al., 2016 ▸; Walsh & Wencewicz, 2013 ▸).

3.2. Model determination and overall structure  

We then performed protein X-ray crystallography in an attempt to obtain snapshots of liganded structures in different states for Hmo and its Y128F mutant. The initial phase problem was solved by the molecular-replacement (MR) method using MDH and FCB2 as search models (Li et al., 2007 ▸; Sukumar et al., 2004 ▸). The solved structure of Hmo was used to generate other ligand-bound structures. The resolutions of the 18 structures ranged from 1.3 to 1.9 Å, all with good R work and R free values. (The diffraction parameters, refinement statistics and PDB codes are summarized in Table 1 ▸.) The structures of Hmo and its mutant with or without ligands all have a single polypeptide chain in the asymmetric unit, while a crystallo­graphic twofold axis generates a dimer. Gel-filtration chromatography and analytical ultracentrifugation analysis (Supplementary Fig. S6) also indicated that dimers are the most biologically relevant state for Hmo. Each monomer is made of a single (β/α)8-barrel domain, known as a TIM barrel, in which the C-terminal loops of the β1, β2 and β8 strands of the barrel constitute the substrate- and cofactor-binding sites [Fig. 3 ▸(a), Supplementary Fig. S7]. One tightly bound FMN cofactor is deeply buried inside the protein, while its redox-active isoalloxazine is accessible to the bulk solvent or substrate (Supplementary Fig. S7). Ternary complexes of the wild type or the Y128F mutant with (S)-p-hydroxy­mandelate or (S)-mandelate (substrates), oxidized mandelate [soaked with (S)-mandelate but oxidized to benzoylformate], p-hydroxy­benzoylformate or benzoylformate (products) or (S)-2-phenylpropionate (an analog/inhibitor) were obtained, in which each ligand fits well to the corresponding electron density [Figs. 3 ▸(a)–3 ▸(c) and 4 ▸(a)–4 ▸(f); Supplementary Figs. S8 and S9]. The physiological substrates other than enantiomers [for example (R)-mandelate], the competitive inhibitors and the products identified here were not anticipated, while these comprehensive complexes are likely to only be obtained in the case of Hmo and its mutants in the α-hydroxyacid oxidase family. This difficulty in and rarity of obtaining such complexes is commonly ascribed to the high substrate-turnover rates in flavin-dependent oxido­reductases (k cat/K m > 105  M −1 s−1; too fast to retain their nascent states) even under flash-soaking/cryogenic cooling, as opposed to those of Hmo and its mutants, which are relatively slower by one order of magnitude (k cat/K m ≃ 104  M −1 s−1).

Figure 3.

Figure 3

Open in a new tab

Liganded structures of Hmo. (a) Superposition of ternary complexes of wild-type Hmo on those of the Y128F mutant shows a low average root-mean-square deviation (r.m.s.d.) of 0.064 Å, where Hmo and the Y128F mutant are colored cyan and green, respectively. There is no significant difference between Hmo (green) and the Y128F mutant (cyan) in terms of the active-site geometry, except that the p-OH group is absent on the phenyl ring of Y128F. (b) When Hmo is bound by (S)-mandelate, Met160 flips away and Arg163 makes an electrostatic association with the carboxylic group of (S)-mandelate. (c) When benzoylformate is bound to Hmo, it adopts double conformations (pro-S or pro-R). Met160 stows back, while Arg163 flips away. The numbers are bond lengths in Å for the designated bonds. Free ligands, FMN and active-site residues are colored cyan, yellow and green, respectively. The 2F o − F c electron-density map is contoured at 2σ.

Figure 4.

Figure 4

Open in a new tab

Ternary-complex structures of the Y128F mutant. (a) The Y128F mutant soaked with benzoylformate, which mainly adopts a pro-R conformation. (b) The Y128F mutant liganded with benzoate, which is the product of the four-electron oxidation reaction catalyzed by the Y128F mutant in the presence of (S)-mandelate. (c) The Y128F mutant liganded with phenylpyruvate, an oxidized product, when soaked with phenyllactate. (d) The Y128F mutant liganded with phenylacetate, which is the product of the four-electron oxidation reaction catalyzed by the Y128F mutant in the presence of 2-­phenyllactate. (e) Y128F bound by the α-hydroxylacid analog 2-hydroxypropanoate. (f) When the Y128F mutant is bound by (R)-mandelate, the structural conformation is similar to that of (S)-mandelate, while the α-OH group points away from His252. The free ligands, FMN and active-site residues are shown in cyan, yellow and green, respectively. The 2F o − F c electron-density map is contoured at 2σ.

3.3. Structural comparison and ligand-binding site  

Superposition of the ternary complexes with (S)-p-hydroxymandelate, (S)-mandelate, benzoylformate or (S)-2-phenylpropionate on the binary complex shows low average root-mean-square deviations (r.m.s.d.s) for 367 Cα backbone atoms, suggesting that there are no significant conformational changes when bound to or lacking a ligand [Fig. 3 ▸(a); Supplementary Fig. S10]. Some local structures do display conformational changes: Met160 near to the ligand entrance may serve as a gatekeeper at the entrance to the ligand-binding tunnel, where the closing/opening of the tunnel synchronizes with the absence/presence of a ligand [Fig. 3 ▸(b)]. Arg163 and Arg255 above and to the right of the isoalloxazine ring interact with the carboxylic end of an α-hydroxyacid through electrostatic forces. An α-ketoacid, however, makes no contact with Arg163, as the guanidino group flips 180° away [Figs. 3 ▸(b) and 3 ▸(c)]. The ligand-binding site is made up of six major residues (Phe24, Ala79, Tyr128, Met160, Arg163, His252 and Arg255) above the si-face (relative to the sp 2 N5) of the isoalloxazine ring [Figs. 3 ▸(a), 3 ▸(b) and 3 ▸(c)], where (S)-p-hydroxymandelate (substrate), (S)-mandelate (substrate) or (S)-2-phenylpropionate (inhibitor) all adopt a ‘V’ conformation with the carboxyl group orthogonal to the plane of the phenyl ring (Supplementary Figs. S8 and S9). Despite being enclosed by hydrophobic residues (Phe24, Ala79 and Tyr128), the cusp of the phenyl ring protrudes towards the funnel-like substrate entrance and is accessible to bulk solvent, allowing some extent of substrate promiscuity, which is consistent with the biochemical analysis [Fig. 3 ▸(a)]. The α-OH group is at low-barrier hydrogen-bond distances to Tyr128 (2.5 Å) and His252 (2.7 Å). Collectively, the α-hydroxyacid is anchored in place by four residues, Tyr128, Arg163, His252 and Arg255, where the α-H points towards N5 of the isoalloxazine ring at a distance of 3.0 Å, in line with the chirality of the reaction [Fig. 3 ▸(b)]. The importance of these residues was examined by site-directed mutagenesis, in which all mutants (R163A, H252F and R255A) except Y128F lost their activity. Moreover, the α-ketoacid and phenyl moieties of the product benzoylformate extend in a planar manner and sit above and in front of the isoalloxazine ring [Fig. 3 ▸(c)].

3.4. Catalytic mechanism  

Tyr128 and His252 in the active site are highly conserved in the family of FMN-dependent α-hydroxyacid oxidases, in which they often serve as a catalytic dyad (Supplementary Fig. S2). The spatial arrangement of the substrate (S)-mandelate and the cofactor FMN in the crystal structures of Hmo favors the HT mechanism, because the distance between the N∊ atom of His252 and α-OH (2.5 Å) is shorter than that between the N∊ atom and C′α (3.0 Å). Crystals soaked with the inhibitors (S)-2-phenylpropionate or (R)-mandelate also show a similar distance (3.1 Å) between N∊ and C′α as that for (S)-mandelate but are devoid of reactivity [Fig. 4 ▸(f), Supplementary Fig. S9]. In this context, His252 is likely to act as the base deprotonating α-OH to form an oxyanion that is stabilized by Tyr128 (Dubois et al., 1990 ▸; Gondry et al., 2001 ▸). Upon collapse of the oxyanion, an α-hydride is expelled to FMN in a hydrophobic chamber at a short distance, fulfilling the reductive half-reaction. FMNox is regenerated by relaying one electron from FMNred to O2, forming a superoxide–FMN semiquinone radical pair, prior to its release as H2O2 in the oxidative half-reaction to complete one round of the catalytic cycle (Fig. 5 ▸; Massey, 1995 ▸; Mattevi, 2006 ▸). While the crystals were soaked with (S)-mandelate, the substrate was converted to benzoylformate. The product unexpectedly adopts two different spatial orientations, pro-S or pro-R, pointing towards or away from the si face of the isoalloxazine ring, respectively [Fig. 3 ▸(c)]. Both the pro-S and pro-R orientations have a similar coplanar conformation without interacting with Arg163. However, when crystals were soaked with benzoylformate the pro-R orientation was almost always observed (Supplementary Fig. S11). We reasoned that the pro-S product is initially formed and that it then converts to the thermodynamically more favorable pro-R orientation. This may be owing to a short distance between the α-keto group of pro-S benzoylformate and p-OH of Tyr128 and N∊ of His252 (2.3 and 2.8 Å, respectively) as a result of the van der Waals repulsion force, thus facilitating the release of benzoylformate (the first two-electron oxidation product) from the binding site [Fig. 3 ▸(c)]. In the Y128F mutant, the efficiency of releasing the product from the active site may be somewhat hampered.

Figure 5.

Figure 5

Open in a new tab

The catalytic cycle of Hmo in the conversion of (S)-mandelate to benzoylformate with the concomitant formation of H2O2.

3.5. Conversion from oxidase to oxidative decarboxylase  

Two mutants, Y128F and H252F, were subjected to a biochemical examination. The latter (H252F) showed no detectable activity, consistent with its designated role, whereas the former (Y128F) showed the activity of an LMO-like enzyme and catalytically converted (S)-mandelate to benzoate [Figs. 1 ▸(d) and 2 ▸(b)]. In detail, no products (benzoate and H2O2) were detected when the wild type (WT) or Y128F mutant was assayed in an anaerobic (no O2) condition containing (S)-mandelate or benzoylformate, suggesting that O2 is the prerequisite electron acceptor in the oxidation reaction. Despite the fact that H2O2 can act as either an oxidant or a reductant (Lopalco et al., 2016 ▸), there was still no product under the same reaction conditions with the addition of H2O2 (2 h, pH 7.3, 2 mM H2O2 with the WT or the Y128F mutant). No product was again detected when benzoylformate was incubated aerobically (with O2) with the WT or the Y128F mutant plus H2O2. As a result, benzoate can only be formed from (S)-mandelate under aerobic conditions, despite benzoylformate and H2O2 being the apparent intermediate and oxidant, respectively. We thus speculated that the intermediate, the oxidant and FMN ought to associate in a well organized manner to allow the second two-electron oxidation to proceed. To prove this, we performed a stable isotope-labeling experiment. Using (S)-mandelate, H2 18O2 and the Y128F mutant under an anaerobic condition, no benzoate was detected. In contrast, 18O-benzoate was detected when the reactions were performed under the same condition with 18O2 in lieu of H2 18O2 [Fig. 2 ▸(c)]; thus, we conclude that free H2O2 is not the effective oxidant.

The conversion rates of (S)-mandelate to benzoylformate or to benzoate by the Y128F mutant were estimated: k cat = 0.05 s−1, K m = 0.13 mM and k cat = 4.5 s−1, K m = 0.15 mM, respectively. The K m values are comparable, suggesting that the substrate-binding affinity of the WT or the Y128F mutant is similar, but the latter specializes in four-electron oxidation. The binding affinity of benzoylformate for Hmo or the Y128F mutant was further determined by ITC. In view of the binding affinity, the discrepancy between the affinity of benzoyl­formate for Hmo (K d = 1.9 mM) and that of benzoylformate for the Y128F mutant (K d = 1.4 mM) is marginal, with the latter being slightly stronger than the former (Supplementary Fig. S5), suggesting that the single mutation has little influence but allows benzoylformate to better interact with the Y128F mutant. The level of H2O2 in reactions with the Y128F mutant is inverse to the level of benzoate, in contrast to those with the WT, in which the level of H2O2 is proportional to that of benzoylformate [Fig. 2 ▸(d)]. R163L, a low-activity mutant, was further assayed against α-(S)-mandelamide [2-(S)-hydroxy-2-phenylethylamide]. This substrate can be recognized and oxidized to the α-ketoamide by the R163L mutant, but not to benzoate [Fig. 2 ▸(e)], indicating that the timing of the decarboxylation step is critical to the four-electron oxidation. In brief, one O atom from O2 is incorporated into benzoate, where the peroxide is not in the form of free H2O2 in solution, but in a well organized manner in which the α-ketoacid and FMNred engage in the oxidative decarboxylation reaction, well beyond the non-ping-pong mechanism described for LMO (Choong & Massey, 1980 ▸; Ghisla & Massey, 1977 ▸; Lockridge et al., 1972 ▸; Walsh et al., 1973 ▸). As revealed above, the pro-R orientation instead of the pro-S is in line with the trajectory of the attack of C4α-peroxide (a Baeyer–Villiger-type reaction) or the nucleophilic sp 3 N5 of FMNred on the α-keto carbonyl C atom of the α-ketoacid, implying that stabilization of the peroxide intermediate and swift reorientation of the products are possible reasons for the conversion of the Y128F mutant from an oxidase to an oxidative decarboxylase [Fig. 4 ▸(a)] (Lyu et al., 2019 ▸).

4. Conclusions  

While flavoproteins, which are omnipresent in all living organisms, have been studied for over a century, many outstanding issues remain unresolved. In this study, structural biology in conjunction with biochemical examinations allowed us to dissect the oxidation reactions carried out by Hmo in steps. In the two-electron oxidation reaction, FMNox is reduced by a hydride directly from an α-hydroxyacid; FMNred is instantly oxidized by O2 to yield H2O2. The collection of multiple liganded snapshots of phases in the active site tangibly supports the HT mechanism and meanwhile accounts for some extent of substrate promiscuity. In the Y128F mutant, the two-electron oxidation reaction is catalytically extended to a four-electron oxidation reaction. The rationale for the reaction is proposed on the basis of the structural information obtained as follows: the absence of the p-OH group in the Y128F mutant substantiates the peroxide anion adduct as a nucleophilic group, and the reorientation of the α-ketoacid in the pro-R configuration above the isoalloxazine ring in the Y128F mutant collaterally provides a better attacking trajectory for nucleophilic FMNred or C4α-peroxide. The Y128F mutant is thereby transformed into a monooxygenase-like enzyme that catalyzes the oxidative decarboxylation reaction, in which the phenolic O atom of Tyr128 in Hmo apparently plays a pivotal role in control of the oxidative cascade through activating and stabilizing reactants and intermediates in addition to facilitating the release of products.

Supplementary Material

Supplementary materials and methods, figures and tables. DOI: 10.1107/S2059798319009574/ag5030sup1.pdf

d-75-00733-sup1.pdf (2MB, pdf)

PDB reference: p-hydroxymandelate oxidase, 5zzp

PDB reference: complex with 4-hydroxy-(S)-mandelate, 5zzq

PDB reference: complex with (S)-mandelate, 5zzr

PDB reference: complex with benzoate, 5zzs

PDB reference: complex with 2-phenylpropanoate, 6a00

PDB reference: complex with benzoylformate, 6a08

PDB reference: Y128F mutant, 6a13

PDB reference: complex with (R)-mandelate, 5zzx

PDB reference: complex with 2-hydroxypropanoate, 5zzy

PDB reference: complex with 2-phenylpropanoate, 6a0d

PDB reference: complex with 2-hydroxy-3-phenylpropanoate, 6a0g

PDB reference: complex with 2-phenylacetate, 6a0m

PDB reference: complex with benzaldehyde, 6a0o

PDB reference: complex with (S)-mandelate, 6a0v

PDB reference: complex with benzoate, 6a0y

PDB reference: complex with phenylpyruvic acid, 6a11

PDB reference: complex with benzoylformate, 6a19

PDB reference: complex with 4-hydroxy-(S)-mandelate, 6a1a

Acknowledgments

Portions of this research were carried out at the National Synchrotron Radiation Research Center (NSRRC), a national user facility supported by MOST of Taiwan, ROC. We thank both NSRRC in Taiwan and SPring-8 in Japan for beamtime allocations at beamlines 13C, 13B, 05A, 15A and 44XU.

Funding Statement

This work was funded by Ministry of Science and Technology, Taiwan grants 102-2311-B-001-028-MY3, 105-2311-B-001-050, and 106-2113-M-001-013-MY2. Academia Sinica grant .

References

  1. Afonine, P. V., Grosse-Kunstleve, R. W., Echols, N., Headd, J. J., Moriarty, N. W., Mustyakimov, M., Terwilliger, T. C., Urzhumtsev, A., Zwart, P. H. & Adams, P. D. (2012). Acta Cryst. D68, 352–367. [DOI] [PMC free article] [PubMed]
  2. Al Toma, R. S., Brieke, C., Cryle, M. J. & Süssmuth, R. D. (2015). Nat. Prod. Rep. 32, 1207–1235. [DOI] [PubMed]
  3. Cao, Y., Han, S., Yu, L., Qian, H. & Chen, J.-Z. (2014). J. Phys. Chem. B, 118, 5406–5417. [DOI] [PubMed]
  4. Chen, Z.-W., Vignaud, C., Jaafar, A., Levy, B., Gueritte, F., Guenard, D., Lederer, F. & Mathews, F. S. (2012). Biochimie, 94, 1172–1179. [DOI] [PubMed]
  5. Choong, Y. S. & Massey, V. (1980). J. Biol. Chem. 255, 8672–8677. [PubMed]
  6. Cunane, L. M., Barton, J. D., Chen, Z.-W., Lê, K. H. D., Amar, D., Lederer, F. & Mathews, F. S. (2005). Biochemistry, 44, 1521–1531. [DOI] [PubMed]
  7. Dijkman, W. P., de Gonzalo, G., Mattevi, A. & Fraaije, M. W. (2013). Appl. Microbiol. Biotechnol. 97, 5177–5188. [DOI] [PubMed]
  8. Dubois, J., Chapman, S. K., Mathews, F. S., Reid, G. A. & Lederer, F. (1990). Biochemistry, 29, 6393–6400. [DOI] [PubMed]
  9. Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. (2010). Acta Cryst. D66, 486–501. [DOI] [PMC free article] [PubMed]
  10. Fagan, R. L. & Palfey, B. A. (2010). Comprehensive Natural Products II: Chemistry and Biology, edited by H.-W. Liu & L. Mander, Vol. 7, pp. 37–113. Kidlington: Elsevier.
  11. Ghisla, S. & Massey, V. (1977). J. Biol. Chem. 252, 6729–6735. [PubMed]
  12. Ghisla, S. & Massey, V. (1989). Eur. J. Biochem. 181, 1–17. [DOI] [PubMed]
  13. Giegel, D. A., Williams, C. H. & Massey, V. (1990). J. Biol. Chem. 265, 6626–6632. [PubMed]
  14. Gillet, N., Ruiz-Pernía, J. J., de la Lande, A., Lévy, B., Lederer, F., Demachy, I. & Moliner, V. (2016). Phys. Chem. Chem. Phys. 18, 15609–15618. [DOI] [PubMed]
  15. Gondry, M., Dubois, J., Terrier, M. & Lederer, F. (2001). Eur. J. Biochem. 268, 4918–4927. [DOI] [PubMed]
  16. Hubbard, B. K., Thomas, M. G. & Walsh, C. T. (2000). Chem. Biol. 7, 931–942. [DOI] [PubMed]
  17. Jones, J. M., Morrell, J. C. & Gould, S. J. (2000). J. Biol. Chem. 275, 12590–12597. [DOI] [PubMed]
  18. Lederer, F., Vignaud, C., North, P. & Bodevin, S. (2016). Biochim. Biophys. Acta, 1864, 1215–1221. [DOI] [PubMed]
  19. Lehoux, I. E. & Mitra, B. (1999). Biochemistry, 38, 5836–5848. [DOI] [PubMed]
  20. Li, S. J., Umena, Y., Yorita, K., Matsuoka, T., Kita, A., Fukui, K. & Morimoto, Y. (2007). Biochem. Biophys. Res. Commun. 358, 1002–1007. [DOI] [PubMed]
  21. Li, T.-L., Choroba, O. W., Charles, E. H., Sandercock, A. M., Williams, D. H. & Spencer, J. B. (2001). Chem. Commun., pp. 1752–1753. [DOI] [PubMed]
  22. Li, T.-L., Huang, F., Haydock, S. F., Mironenko, T., Leadlay, P. F. & Spencer, J. B. (2004). Chem. Biol. 11, 107–119. [DOI] [PubMed]
  23. Lindqvist, Y. (1989). J. Mol. Biol. 209, 151–166. [DOI] [PubMed]
  24. Lockridge, O., Massey, V. & Sullivan, P. A. (1972). J. Biol. Chem. 247, 8097–8106. [PubMed]
  25. Lopalco, A., Dalwadi, G., Niu, S., Schowen, R. L., Douglas, J. & Stella, V. J. (2016). J. Pharm. Sci. 105, 705–713. [DOI] [PMC free article] [PubMed]
  26. Lyu, S.-Y., Lin, K.-H., Yeh, H.-W., Li, Y.-S., Huang, C.-H., Wang, Y.-L., Shih, H.-W., Hsu, N.-S., Wu, C.-J. & Li, T.-L. (2019). Submitted.
  27. Maeda-Yorita, K., Aki, K., Sagai, H., Misaki, H. & Massey, V. (1995). Biochimie, 77, 631–642. [DOI] [PubMed]
  28. Massey, V. (1995). FASEB J. 9, 473–475. [DOI] [PubMed]
  29. Mattevi, A. (2006). Trends Biochem. Sci. 31, 276–283. [DOI] [PubMed]
  30. McCafferty, D. G., Cudic, P., Frankel, B. A., Barkallah, S., Kruger, R. G. & Li, W. (2002). Biopolymers, 66, 261–284. [DOI] [PubMed]
  31. McCoy, A. J., Grosse-Kunstleve, R. W., Adams, P. D., Winn, M. D., Storoni, L. C. & Read, R. J. (2007). J. Appl. Cryst. 40, 658–674. [DOI] [PMC free article] [PubMed]
  32. Murray, M. S., Holmes, R. P. & Lowther, W. T. (2008). Biochemistry, 47, 2439–2449. [DOI] [PMC free article] [PubMed]
  33. Murshudov, G. N., Skubák, P., Lebedev, A. A., Pannu, N. S., Steiner, R. A., Nicholls, R. A., Winn, M. D., Long, F. & Vagin, A. A. (2011). Acta Cryst. D67, 355–367. [DOI] [PMC free article] [PubMed]
  34. Otwinowski, Z. & Minor, W. (1997). Methods Enzymol. 276, 307–326. [DOI] [PubMed]
  35. Perham, R. N., Jones, D. D., Chauhan, H. J. & Howard, M. J. (2002). Biochem. Soc. Trans. 30, 47–51. [DOI] [PubMed]
  36. Petsko, G. A., Kenyon, G. L., Gerlt, J. A., Ringe, D. & Kozarich, J. W. (1993). Trends Biochem. Sci. 18, 372–376. [DOI] [PubMed]
  37. Romero, E., Gómez Castellanos, J. R., Gadda, G., Fraaije, M. W. & Mattevi, A. (2018). Chem. Rev. 118, 1742–1769. [DOI] [PubMed]
  38. Sukumar, N., Dewanti, A. R., Mitra, B. & Mathews, F. S. (2004). J. Biol. Chem. 279, 3749–3757. [DOI] [PubMed]
  39. Tsou, A. Y., Ransom, S. C., Gerlt, J. A., Buechter, D. D., Babbitt, P. C. & Kenyon, G. L. (1990). Biochemistry, 29, 9856–9862. [DOI] [PubMed]
  40. Wageningen, A. A. van, Kirkpatrick, P. N., Williams, D. H., Harris, B. R., Kershaw, J. K., Lennard, N. J., Jones, M., Jones, S. J. & Solenberg, P. J. (1998). Chem. Biol. 5, 155–162. [DOI] [PubMed]
  41. Walker, S., Chen, L., Hu, Y., Rew, Y., Shin, D. & Boger, D. L. (2005). Chem. Rev. 105, 449–476. [DOI] [PubMed]
  42. Walsh, C., Lockridge, O., Massey, V. & Abeles, R. (1973). J. Biol. Chem. 248, 7049–7054. [PubMed]
  43. Walsh, C. T. & Wencewicz, T. A. (2013). Nat. Prod. Rep. 30, 175–200. [DOI] [PMC free article] [PubMed]
  44. Winn, M. D., Ballard, C. C., Cowtan, K. D., Dodson, E. J., Emsley, P., Evans, P. R., Keegan, R. M., Krissinel, E. B., Leslie, A. G. W., McCoy, A., McNicholas, S. J., Murshudov, G. N., Pannu, N. S., Potterton, E. A., Powell, H. R., Read, R. J., Vagin, A. & Wilson, K. S. (2011). Acta Cryst. D67, 235–242. [DOI] [PMC free article] [PubMed]
  45. Xia, Z.-X. & Mathews, F. S. (1990). J. Mol. Biol. 212, 837–863. [DOI] [PubMed]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary materials and methods, figures and tables. DOI: 10.1107/S2059798319009574/ag5030sup1.pdf

d-75-00733-sup1.pdf (2MB, pdf)

PDB reference: p-hydroxymandelate oxidase, 5zzp

PDB reference: complex with 4-hydroxy-(S)-mandelate, 5zzq

PDB reference: complex with (S)-mandelate, 5zzr

PDB reference: complex with benzoate, 5zzs

PDB reference: complex with 2-phenylpropanoate, 6a00

PDB reference: complex with benzoylformate, 6a08

PDB reference: Y128F mutant, 6a13

PDB reference: complex with (R)-mandelate, 5zzx

PDB reference: complex with 2-hydroxypropanoate, 5zzy

PDB reference: complex with 2-phenylpropanoate, 6a0d

PDB reference: complex with 2-hydroxy-3-phenylpropanoate, 6a0g

PDB reference: complex with 2-phenylacetate, 6a0m

PDB reference: complex with benzaldehyde, 6a0o

PDB reference: complex with (S)-mandelate, 6a0v

PDB reference: complex with benzoate, 6a0y

PDB reference: complex with phenylpyruvic acid, 6a11

PDB reference: complex with benzoylformate, 6a19

PDB reference: complex with 4-hydroxy-(S)-mandelate, 6a1a

Articles from Acta Crystallographica. Section D, Structural Biology are provided here courtesy of International Union of Crystallography

RESOURCES