Prostaglandin H synthase: resolved and unresolved mechanistic issues

Abstract

The cyclooxygenase and peroxidase activities of prostaglandin H synthase (PGHS)-1 and -2 have complex kinetics, with the cyclooxygenase exhibiting feedback activation by product peroxide and irreversible self-inactivation, and the peroxidase undergoing an independent self-inactivation process. The mechanistic bases for these complex, non-linear steady-state kinetics have been gradually elucidated by a combination of structure/function, spectroscopic and transient kinetic analyses. It is now apparent that most aspects of PGHS-1 and -2 catalysis can be accounted for by a branched chain radical mechanism involving a classic heme-based peroxidase cycle and a radical-based cyclooxygenase cycle. The two cycles are linked by the Tyr385 radical, which originates from an oxidized peroxidase intermediate and begins the cyclooxygenase cycle by abstracting a hydrogen atom from the fatty acid substrate. Peroxidase cycle intermediates have been well characterized, and peroxidase self-inactivation has been kinetically linked to a damaging side reaction involving the oxyferryl heme oxidant in an intermediate that also contains the Tyr385 radical. The cyclooxygenase cycle intermediates are poorly characterized, with the exception of the Tyr385 radical and the initial arachidonate radical, which has a pentadiene structure involving C11-C15 of the fatty acid. Oxygen isotope effect studies suggest that formation of the arachidonate radical is reversible, a conclusion consistent with electron paramagnetic resonance spectroscopic observations, radical trapping by NO, and thermodynamic calculations, although moderate isotope selectivity was found for the H-abstraction step as well. Reaction with peroxide also produces an alternate radical at Tyr504 that is linked to cyclooxygenase activation efficiency and may serve as a reservoir of oxidizing equivalent. The interconversions among radicals on Tyr385, on Tyr504, and on arachidonate, and their relationships to regulation and inactivation of the cyclooxygenase, are still under active investigation for both PGHS isozymes.

Scheme 1

A hypothetical model summarizing the interconversions of the various tyrosyl radical species and the AA radical in PGHS-1 and -2. The EPR signatures are indicated for each of the tyrosyl radicals, along with the assigned location of the radical; the AA radical (AA•) is the pentadienyl species with a 7-line EPR signal. Parallel reactions for formation of the Tyr385 and Tyr504 radicals from Intermediate I are indicated by the yellow arrows at the top. AA reacts with the Tyr385 radical to form the AA radical (AA•); in PGHS-1, AA can also bind to the Tyr385 radical (without H-abstraction) to form the NS1a/NS1c radical, thought to be located on Tyr504, Tyr348, or Tyr385 (conformation differs from that in WD1). O₂ reacts with the AA• to eventually form PGG₂. In trapping experiments, NO reacts with the AA• to form oximes. The specificity of individual steps to one isozyme or the other is indicated by color coding. Cyclooxygenase inactivation steps are shown as dashed red arrows.

Figure 1

Reaction of PGHS with AA showing the cyclooxygenase and peroxidase steps. Changes in the atoms derived from molecular oxygen are highlighted with dashed lines.

Figure 2

Structure of AA bound to PGHS-1 from crystallographic data (1DIY) [6]. Heme was replaced by cobalt protoporphyrin IX (shown in green); AA (backbone shown in cyan) interacts electrostatically with Arg120, with C13 of the fatty acid adjacent to Tyr385; and Ser530, the target of aspirin, is nearby. The locations of the membrane anchor helices and the trypsin digestion site, Arg277, are also indicated.

Figure 3

Time courses for peroxidase (left) and cyclooxygenase reactions of PGHS-1. The peroxidase reaction with HOOH was monitored by A₄₃₆ increases due to oxidation of the reducing cosubstrate, guaiacol. The cyclooxygenase reaction with AA was followed by measuring oxygen uptake with a polarographic electrode [143]. The peak velocity in each reaction is indicated by a dashed line.

Figure 4

Current PGHS peroxidase mechanism. The generally accepted mechanism [33] is a hybrid of the mechanisms of horseradish peroxidase and cytochrome c peroxidase. The formal oxidation state of the heme iron is indicated. ROOH and ROH, hydroperoxide and corresponding alcohol; PPIX and PPIX•⁺, protoporphyrin IX and its cation radical; e-, reducing equivalent from peroxidase cosubstrate; Y•, tyrosyl radical.

Figure 5

Structural model of key residues in PGHS peroxidase catalysis, based on crystallographic data for PGHS-1 containing MnPPIX (2AYL) [53]. The protoporphyrin is shown in yellow, with Mn as an orange sphere. The structured water H-bonded between Tyr504 and His388 is shown as a cyan sphere. Distances are indicated in Å.

Figure 6

X-band EPR spectra of oPGHS-1 before (red) and after (blue) reaction with EtOOH. The resting enzyme has prominent signals from high-spin ferric heme (HS) and low-spin ferric heme (LS), along with a small signal from non-specific ferric iron (NI). The heme signals are decreased in the peroxide-reacted PGHS-1, replaced by the tyrosyl radical signal (RAD) with a g value of ~2. Inset: expanded view of radical signal showing the hyperfine features. The resting PGHS-1 spectrum is essentially featureless in the g = 2 region when recorded at the low modulation amplitude used for peroxide-treated PGHS-1. Reproduced with permission from Ref. [21] (Copyright 2000 Elsevier Science Inc.).

Figure 7

The “branched-chain” reaction mechanism linking PGHS peroxidase and cyclooxygenase. The model originally proposed by Ruf and colleagues [33, 34] is adapted here to indicate that the PGHS peroxidase (POX) might be considered a “receptor” for peroxide “ligand”, leading to formation of the “transducer”, the Tyr385 radical, and thus activating the “effector”, cyclooxygenase catalysis. An imaginary boundary between “receptor” and “effector” is indicated by a dashed line.

Figure 8

X-band EPR signals of peroxide-induced tyrosyl radicals observed in oPGHS-1 and hPGHS-2. Assignments for individual signals are shown in the table at right.

Figure 9

Geometry of the radical tyrosyl side chain. Left: illustration of the localization of the unpaired electron at C1, C3 and C5, and showing the key dihedral angle, θ, and the direction of the three principal g components. Right: numbering of carbon atoms.

Figure 10

Comparison of HFEPR and x-band EPR spectra of WD1, WS1, and NS1a radical signals from oPGHS-1 with spectra of the tyrosyl radical in ribonucleotide reductase. HFEPR spectra were recorded at 190 and 285 GHz, and x-band spectra (insets) at 9 GHz. The WD1 spectra are superimposed on the ribonucleotide reductase Tyr• spectra in the lower right panel. Reproduced with permission from Ref. [64] (Copyright 2000 American Chemical Society).

Figure 11

Components of PGHS tyrosyl radical dynamics. Structure of oPGHS-1 (1DIY; [6]) illustrating the position of Tyr385, which is H-bonded to Tyr348, Tyr504, Tyr148, Tyr404, the potential alternate tyrosyl radical site, and bound AA, which has C13 shown in green. Potential conformational changes in the Tyr385 sidechain are indicated by arrows. Heme is replaced by CoPPIX in this structure.

Figure 12

EPR spectra of hPGHS-1 and -2 mutants supporting alternate radical formation at Tyr504. Samples of hPGHS-2 constructs (A) and hPGHS-1 constructs (B) [68] were reacted with EtOOH and manually freeze trapped before the EPR spectra were recorded [68, 69]. Reproduced with permission from Ref. [69] (Copyright 2009 Elsevier Science Inc.) and from Ref. [68] (Copyright 2004 American Chemical Society).

Figure 13

Evidence for kinetic competence of WD1 tyrosyl radical in oPGHS-1. Correlation of oxidized heme intermediates (decrease in A₄₁₀ and increase in A₅₈₀), WD1 (doublet) tyrosyl radical, and cyclooxygenase product (PGG₂ + PGH₂, total eicosanoids) during aerobic reaction of oPGHS-1 with AA [60]. Reproduced with permission from Ref. [60] (Copyright 1992 American Society for Biochemistry and Molecular Biology).

Figure 14

Evidence for kinetic competence of WD1 tyrosyl radical in oPGHS-1. Top panel: Rapid freeze quench EPR kinetic measurements showing rapid formation of WD1 in reactions of oPGHS-1 with EtOOH [57] and the time-dependent spectral transition to WS1. Bottom: Kinetics of tyrosyl radical concentration (spin/heme) and ferryl heme levels (A₅₂₄). Reproduced with permission from Ref. [57] (Copyright 1999 American Society for Biochemistry and Molecular Biology).

Figure 15

Hypothetical mechanism for cyclooxygenase catalysis, modified from the original proposal by Hamberg and Samuelson [80] to include the role of the Tyr385 radical (Y•) in generation of the first AA radical intermediate (AA•) and to reflect the possible reversibility of the initial steps (blue arrows). Anaerobic conditions block at step 2.

Figure 16

EPR spectra supporting generation of the first AA radical intermediate by PGHS-2 tyrosyl radical (step 1 in Fig. 15). PGHS-2 was mixed sequentially with EtOOH and then AA under anaerobic conditions before freeze-quenching and EPR analysis [82]. The spectrum obtained before addition of AA (a) shows the WS2 signal reflecting a mixture of Tyr385 and Tyr504 radicals [68]. The spectra obtained at shorter and longer times after addition of AA (b, c) show the 7-line hyperfine splitting pattern consistent with an AA pentadienyl radical [82]. Thawing and adding oxygen to these samples regenerated the WS2 tyrosyl radical EPR (b′ and c′). Radical intensities are given in spin/heme. Reproduced with permission from Ref. [82] (Copyright 1998 American Society for Biochemistry and Molecular Biology).

Figure 17

Structure of the initial AA radical in cyclooxygenase catalysis. Panel a: Conformation of C8-C16 of the pentadienyl AA radical deduced from EPR characterization and additional constraints from the geometry of the C9-C11 endoperoxide and the C8-C12 crosslink [82]. The locations of the six strongly-interacting protons are highlighted, along with the spin distribution and the key dihedral angles at C10 and C16. Panel b: Comparison of the AA radical conformation deduced from EPR analysis (center) with the protein-bound structures of AA (left, 1DIY) and PGG₂ (right, 1DDX). C1-C7 and C17-C20 of the AA radical were arranged manually to approximate their positions in the AA and PGG₂ structures.

Figure 17

Structure of the initial AA radical in cyclooxygenase catalysis. Panel a: Conformation of C8-C16 of the pentadienyl AA radical deduced from EPR characterization and additional constraints from the geometry of the C9-C11 endoperoxide and the C8-C12 crosslink [82]. The locations of the six strongly-interacting protons are highlighted, along with the spin distribution and the key dihedral angles at C10 and C16. Panel b: Comparison of the AA radical conformation deduced from EPR analysis (center) with the protein-bound structures of AA (left, 1DIY) and PGG₂ (right, 1DDX). C1-C7 and C17-C20 of the AA radical were arranged manually to approximate their positions in the AA and PGG₂ structures.

Figure 18

EPR spectra of AA radicals produced by hPGHS-2. Peroxide-treated hPGHS-2 was reacted anaerobically with unlabeled AA or with the indicated deuterium-substituted AA. Spectral simulations using the EPR parameter set in Ref. [82], adjusted for deuterium substitution, are shown in dashed lines. Pink dots in the structures indicate the deuterated sites, and the hyperfine splitting patterns are indicated at right.

Figure 19

Calculated free energy landscape of the cyclooxygenase catalysis in PGHS based on hybrid density functional theory (AM1/UB3LYP) [96]. Potential rate-limiting steps are indicated by red arrows and reverse steps with high activation barriers are indicated by green arrows. Reproduced with permission from Ref. [96] (Copyright 2003 American Chemical Society).

Figure 20

Sequential stopped-flow device used to analyze PGHS peroxidase and cyclooxygenase inactivation kinetics. The “substrate” syringe for the second stage reaction holds excess H₂O₂+guaiacol (to assay surviving peroxidase activity) or excess 11,14-eicosadienoic acid (to assay surviving cyclooxygenase activity).

Figure 21

Effect of peroxide concentration on oPGHS-1 peroxidase self-inactivation kinetics. Sequential stopped-flow experiments with 7.8-250 μM EtOOH in the first stage used the device shown in Fig. 20. Panel A: Second-stage time courses of surviving peroxidase activity (monitored by oxidation of guaiacol at 436 nm) are shown for the indicated durations of oPGHS-1 first-stage incubation with 250 μM EtOOH. The surviving peroxidase activity was quantified from the maximal slope of each time course. Panel B: Surviving peroxidase activities measured in the second-stage reaction are plotted as a function of the first-stage aging time with each of the indicated levels of EtOOH. First order rate constants were estimated by fitting the data to an exponential equation. Reproduced with permission from Ref. [121] (Copyright 1999 American Society for Biochemistry and Molecular Biology).

Figure 22

Effects of peroxide structure and oPGHS-1 concentration on peroxidase self-inactivation kinetics. The first order rate constants determined for peroxidase self inactivation are shown as a function of peroxide concentration for HOOH (A), EtOOH (B), PPHP (C) and 15-HPETE (D). Inactivation kinetics were measured at several levels of oPGHS-1 (enzyme concentrations are indicated alongside the symbols) for each peroxide. Adapted with permission from Ref. [121] (Copyright 1999 American Society for Biochemistry and Molecular Biology).. Symbols refer to measured rates at different enzyme concentrations.

Figure 23

Hypothetical peroxidase inactivation mechanism deduced from sequential stopped-flow studies. Two new spectroscopic species derived from Intermediate II (“Intermediate III” and “Terminal Compound”) were proposed for the process of irreversible inactivation. Note that Intermediate II contains two potentially damaging oxidants, tyrosyl radical and ferryl heme (highlighted by boxes).

Figure 24

Cosubstrate effects on peroxide-induced inactivation of peroxidase and cyclooxygenase activities in oPGHS-1. Sequential stopped-flow experiments were used to monitor the decay of peroxidase and cyclooxygenase activities during incubation with peroxide (60 μM EtOOH) in the presence or absence of cosubstrate (0.50 mM guaiacol). A one-phase fit was used for decay with guaiacol present and a two-phase fit was used for decay without guaiacol. Reproduced with permission from Ref. [122] (Copyright 2003 American Chemical Society).

Figure 25

Comparison of decay kinetics for tyrosyl radical (Tyr•), oxyferryl heme (A₄₂₈) and peroxidase activity (POX) during reaction of 30 μM oPGHS-1 with 20 equivalents of either EtOOH (top; Ref.[73]) or 15-HPETE (bottom). Reproduced with permission from Ref. [73] (Copyright 2007 American Chemical Society.