A Mechanistic Study of Self-inactivation of the Peroxidase
Activity in Prostaglandin H Synthase-1*
Gang
Wu
,
Chunhong
Wei
,
Richard J.
Kulmacz
,
Yoichi
Osawa§, and
Ah-lim
Tsai
¶
From the
Division of Hematology, Department of
Internal Medicine, University of Texas Health Science Center,
Houston, Texas 77030 and the § Department of Pharmacology,
University of Michigan Medical School,
Ann Arbor, Michigan 48109-0632
 |
ABSTRACT |
Prostaglandin H synthase (PGHS) is a
self-activating and self-inactivating enzyme. Both the peroxidase and
cyclooxygenase activities have a limited number of catalytic turnovers.
Sequential stopped-flow measurements were used to analyze the kinetics
of PGHS-1 peroxidase self-inactivation during reaction with several different hydroperoxides. The inactivation followed single exponential kinetics, with a first-order rate constant of 0.2-0.5
s
1 at 24 °C. This rate was independent of the
peroxide species and concentration used, strongly suggesting that the
self-inactivation process originates after formation of Compound I and
probably with Intermediate II, which contains an oxyferryl heme and a
tyrosyl radical. Kinetic scan and rapid scan experiments were used to monitor the heme changes during the inactivation process. The results
from both experiments converged to a simple, linear, two-step mechanism
in which Intermediate II is first converted in a faster step (0.5-2
s
1) to a new compound, Intermediate III, which undergoes
a subsequent slower (0.01-0.05 s
1) transition to a
terminal species. Rapid-quench and high pressure liquid chromatography
analysis indicated that Intermediate III likely retains an intact heme
group that is not covalently linked with the PGHS-1 protein.
 |
INTRODUCTION |
Prostaglandin H synthase
(PGHS)1 (1) catalyzes the
first committed step in the biosynthesis of many important
prostanoids. PGHS exhibits two enzymatic activities as follows: a
cyclooxygenase activity that converts arachidonic acid to
PGG2 and a peroxidase activity that transforms
PGG2 to PGH2. Several oxidized reaction intermediates have been identified (1-4). PGHS first interacts with a
peroxide substrate to generate Intermediate I, equivalent to Compound I
of horseradish peroxidase. Intermediate I then converts to Intermediate
II, equivalent to Complex ES of cytochrome c
peroxidase, through an intramolecular electron transfer from a tyrosine
residue to the oxidized porphyrin. The transiently formed tyrosyl
radical present in Intermediate II has been demonstrated to be capable of oxidizing arachidonic acid to generate an arachidonic acid radical
and beginning cyclooxygenase catalysis in both PGHS isoforms (5, 6).
X-ray crystallographic data revealed that the presumed site of the
tyrosyl radical, Tyr-385 in PGHS-1 (Tyr-371 in PGHS-2), is located
between the heme- and arachidonate-binding sites, well positioned to
serve a role in coupling the two enzyme activities (7-9). These recent
crystallographic findings substantiate the original branched-chain
mechanism (2, 3) which proposed that a tyrosyl radical generated in the
peroxidase cycle leads to production of PGG2 as long as
arachidonate is present.
One important fundamental limit on prostaglandin synthesis is the
characteristic self-inactivation of PGHS (10). A typical reaction
kinetics profile for PGHS with arachidonate, monitored by oxygen
uptake, shows an initial burst, reaches a maximum velocity, and
gradually decreases to zero (10). This fall in cyclooxygenase activity
is not due to exhaustion of substrate or product inhibition but is
rather a consequence of suicide inactivation (10-13). Each PGHS
molecule thus exhibits only a limited number of turnovers before the
cyclooxygenase activity disappears. With purified PGHS, this number of
turnovers can be as high as 1300 and as low as 10 depending on the
concentration of reductants and other factors (14, 15).
The detailed mechanism of the self-inactivation process has not been
thoroughly characterized. Degradation of both peroxidase and
cyclooxygenase activities have been observed during the
self-inactivation process, as evidenced by decay of enzyme activities
(15-17) and heme spectral changes (18). Based on the protective effect
of various reducing cosubstrates against self-inactivation in both enzyme activities (15, 19-22), a consensus has emerged that certain active radical intermediates generated in peroxidase or cyclooxygenase catalysis lead to irreversible loss of enzyme activity. However, a
direct linkage between a particular radical species and
self-inactivation has yet to be established. The present study
concentrates on peroxidase self-inactivation in the absence of
exogenous reducing substrate to simplify the interpretations. The
results provide convincing evidence that the self-inactivation step
occurs subsequent to formation of Intermediate II and is thus
independent of peroxide structure and concentration.
 |
EXPERIMENTAL PROCEDURES |
Hydrogen peroxide, guaiacol, and hemin were purchased from
Sigma. Arachidonic acid was from Nu Chek Prep (Elysian, MN). EtOOH was
purchased as a 5% aqueous solution from Polysciences Inc. (Warrington,
PA) or from Accurate Chemical and Scientific Corp. (Westbury, NY) as a
10% solution. PPHP was the product of Cayman Chemical Co. (Ann Arbor,
MI). Chloroperbenzoic acid was from Aldrich and was recrystallized in
water. 15-HPETE was prepared according to Graff et al. (23)
using arachidonate and soybean lipoxygenase. The purity of 15-HPETE was
assessed chromatographically and its concentration quantified from the
oxidation of TMPD catalyzed by excess PGHS-1 as described previously
(17). Octyl-
-D-glucopyranoside was purchased from
Calbiochem or Amresco (Solon, OH). Tween 20 was purchased from Pierce
or Bio-Rad.
PGHS was prepared as the apoenzyme from ram seminal vesicles as
described previously, with 5 mM glutathione included in the isoelectric focusing step (4, 24). PGHS holoenzyme was formed by adding
heme to the apoenzyme. Unbound heme was removed by treatment with DE52
cellulose gel (Whatman, UK). The concentration of PGHS holoenzyme was
based on the concentration of incorporated heme determined from the
absorbance at 411 nm (165 mM
1
cm
1).
Cyclooxygenase activity was assayed polarographically at 30 °C as
described previously (24).
Measurement of Time-dependent Decay of Peroxidase
Activity--
These experiments were conducted as two-stage reactions
using a Bio-SEQUENTIAL DX-18MV stopped-flow instrument (Applied
Photophysics, Leatherhead, UK). PGHS holoenzyme was first mixed with a
specific level of peroxide and aged for a defined length of time (100 ms to 10 s). This enzyme/peroxide reaction mixture was then mixed in the second stage with a solution containing guaiacol and hydrogen peroxide. Oxidation of guaiacol (monitored from absorbance changes at
436 nm) in the second stage was used to quantify the surviving peroxidase activity, with the peroxidase velocity calculated from the
slope of the initial linear region of the
A436 versus time traces. We chose
to use the initial velocity and not the final extent of the guaiacol
oxidation to quantify the residual peroxidase activity because
secondary processes, including heme bleaching, complicate the later
part of the reaction. In control experiments, carryover of peroxide
from the first stage reaction was evaluated by including EtOOH or
15-HPETE in the guaiacol/H2O2 solution used in
the second stage reaction and found to have negligible effect on the
observed peroxidase velocity (data not shown).
Kinetics of PGHS Heme Spectral Changes Induced by
Peroxide--
Measurements were performed on the Bio-Sequential
stopped-flow instrument in the kinetic scan-spectral reconstruction
mode or on an Olis RSM-1000 rapid-scan stopped-flow instrument (On-Line Instrument Systems, Inc., Bogart, GA). The former approach was used to
acquire data over shorter wavelength ranges (typically, 390-430 nm in
2-nm increments) and at lower enzyme concentrations. Kinetic scan mode
data obtained at serial wavelengths were first processed by the
singular value decomposition (SVD) method (4) and then fitted to
kinetic models using the Glint analysis package to deconvolute spectral
intermediates. The Olis rapid-scan instrument was used principally to
obtain spectral data in the visible region, where a more concentrated
sample is required. Similar SVD analysis and global fitting routines
were used to analyze data obtained with the Olis instrument to resolve
dominant spectral intermediates and to determine the associated
reaction rate constants. A set of published kinetic data and a
mechanistic model for the PGHS peroxide reaction containing three
spectral species (4) was used to validate the reliability of the
SVD/global fitting software supplied with the Applied Photophysics and
Olis instruments and achieved an excellent agreement.
Examination of Heme Modification by Chemical Quench and HPLC
Analysis--
Covalent changes in the heme during the peroxide-induced
self-inactivation were examined by a combination of rapid quench and
HPLC analysis. PGHS (5-10 µM) was reacted with 10 eq of
EtOOH for a defined period and then quenched with solvent (60%
acetonitrile and 1.2% trifluoroacetic acid) at 24 °C in an Update
Instrument System 1000 Chemical/Freeze Quench Apparatus (Madison, WI).
The final concentrations of the reactants and quenching solvent after the two-stage mixing procedure were one-third the originals.
Chemical-quenched samples were stored on Dry Ice until HPLC analysis.
HPLC was performed with a Waters Model 600S controller and model 996 photodiode array detector and a Vydac C4 column (5 µm;
0.21 × 15 cm) equilibrated with Solvent A (0.1% trifluoroacetic
acid) at a flow rate of 0.3 ml/min. A gradient was run to 75% Solvent
B (0.1% trifluoroacetic acid in acetonitrile) over 30 min and then to
100% Solvent B over the next 5 min. The absorbance was monitored at
220 and 400 nm to detect protein and heme species, respectively.
Control experiments with sperm whale myoglobin and hydrogen peroxide
confirmed efficient trapping of the reaction with this solvent
quenching system (data not shown).
Computer Modeling--
The SCoP program (Simulation Resources
Inc., Redlands, CA) was also used for kinetic simulations and for
fitting of single wavelength kinetic data to the mechanistic model
described below (4). The SCoP program allows testing of more
complicated mechanistic models and direct evaluation of each dependent
variable, providing information regarding the sensitivity of the
overall kinetics to the value of each parameter. The equations in the
SCoP program were derived from the mechanism in
Scheme 1 and are shown in
Table I. The values of
k1 and k2 were from Ref.
4. Values for the floating parameters k7 and
k8 were obtained from spectral deconvolution of
kinetic scan and global fitting which will be detailed under "Results." The rates of the reduction steps
(k3, k4,
k5, and k6) fall in the
range commonly observed with PGHS peroxidase (19-22). No spectral
distinction was made between Fe(IV) and Fe(IV)Tyr· intermediates
or between Fe(III)Tyr· and Fe(III) species. The rate of
reduction of tyrosine radical in species Fe(IV)Tyr· was assumed
to be the same as that in Fe(III)Tyr·.
 |
RESULTS |
Kinetics of PGHS Peroxidase Inactivation during Reaction with
EtOOH--
PGHS was reacted with 250 µM EtOOH for
various times, and the surviving peroxidase activity was monitored by
H2O2-dependent oxidation of
guaiacol in a secondary reaction, as described under "Experimental
Procedures" (Fig. 1A). The
initial velocity in the second stage reactions (with
H2O2/guaiacol) is plotted as a function of the
duration of the first stage reaction (with EtOOH) in Fig. 1B, along with corresponding data involving first stage
reactions with other EtOOH levels. The surviving peroxidase activity
(measured in the second stage reaction) followed essentially the same
single exponential decay kinetics for all EtOOH levels between
7.8 to 250 µM (Fig. 1B), with a first-order
rate constant of ~0.4 s
1 at 24 °C.

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Fig. 1.
Peroxidase self-inactivation kinetics during
reaction with EtOOH. A, kinetics of
H2O2-dependent guaiacol oxidation
in the second stage reaction. PGHS (0.5 µM) was reacted
in the first stage with 250 µM EtOOH in 0.1 M
potassium phosphate, pH 7.2, with 10% glycerol, 0.1%
octyl- -D-glucopyranoside and 0.1% Tween 20 for the
indicated period and then reacted in the second stage with 10 mM H2O2 and 10 mM
guaiacol in 0.1 M Tris, pH 8.0, containing 2 mM
EDTA. B, the surviving peroxidase activity for each length
of first stage reaction with 250 µM EtOOH was calculated
from the initial slope of the A436
versus time trace for the second stage reactions in
A and plotted as a function of the first stage time, along
with similar data from reactions with other EtOOH levels. Single
exponential fits were used to determine the value of
kobs for decay of activity in each data
set.
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|
Effects of Peroxide Structure and Enzyme Level on Kinetics of PGHS
Peroxidase Inactivation--
Kinetic measurements of the peroxidase
self-inactivation were extended to other peroxide substrates, including
H2O2, 15-HPETE, and PPHP, at PGHS levels of
0.2-1.3 µM (Fig. 2). In
each instance, the surviving peroxidase activity was determined by a
second stage reaction with H2O2 and guaiacol,
as described above for the experiments in Fig. 1A. In each
case, the decay in peroxidase activity was found to follow first-order
kinetics as the first stage reaction time was increased (data not
shown). The rates of self-inactivation were calculated by fitting the
data to single-exponential equations, as in Fig. 1B. From
the data in Fig. 2 it is clear that the rate of peroxidase
self-inactivation was essentially independent of peroxide structure and
peroxide concentration for the four peroxides examined, with
rates of 0.20-0.44 s
1 for H2O2,
0.22-0.54 s
1 for EtOOH, 0.22-0.46 s
1 for
PPHP, and 0.1-0.5 s
1 for 15-HPETE. The overall average
self-inactivation rate for the data in Fig. 2 was 0.40 s
1. It is also apparent from the data in Fig. 2
that the rate of peroxidase self-inactivation was insensitive to the
PGHS level over the 0.15-1.5 µM range.

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Fig. 2.
Effects of peroxide concentration and
structure on the rate of peroxidase self-inactivation. Peroxidase
inactivation kinetics were determined using the procedure described in
the legend to Fig. 1 at various levels of four different
hydroperoxides: H2O2 (A), EtOOH
(B), PPHP (C), and 15-HPETE (D).
Inactivation kinetics were evaluated with three or four different
PGHS-1 concentrations for each hydroperoxide, as indicated at the
top of each panel.
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|
PGHS Heme Spectral Intermediates Generated during Peroxidase
Self-inactivation--
Optical spectral changes in the PGHS heme
center were monitored during self-inactivation (Fig.
3). Reconstructed spectra after SVD
analysis from 390 to 430 nm for reaction between 1 µM
PGHS-1 and 30 µM EtOOH are shown in Fig. 3A.
Resting enzyme had a Soret peak near 410 nm (spectrum a).
Early in the reaction with EtOOH (0-0.5 s), the intensity of the Soret
peak decreased and shifted to longer wavelengths, consistent with rapid
sequential formation of Intermediates I and II. Subsequent spectra
obtained after 0.5 s of reaction (thin lines in Fig.
3A) were consistent with a single transition. Spectral
changes in this reaction period were highlighted by a maximal increase
near 392 nm and a maximal decrease near 420 nm, with an isosbestic
point at 405-406 nm. An additional, slower transition (dashed
lines in Fig. 3A) followed, finally yielding a spectrum
characterized by a peak at 405 nm with a low absorption coefficient
(lower heavy line in Fig. 3A). These spectral data after the first 50 ms of reaction were further analyzed by the
global fitting package, Glint, to obtain the spectra of the major
components present in this part of the reaction. A simple mechanism,
represented by Reaction 1, was used for the fitting, because the SVD
analysis indicated three dominant species were present in that time
range.
|
(Reaction 1)
|
The three prominent spectral intermediates resulting from this fit
are shown in Fig. 3B, together with the spectrum of the resting enzyme. The first intermediate considered here (A in
Reaction 1), with a peak at 413-414 nm (spectrum b), is
very similar to published spectra of PGHS-1 Intermediate II, containing
an oxyferryl heme and a tyrosyl radical (1-4). The second intermediate
observed (B in Reaction 1), called Intermediate III,
exhibited a peak at 403-404 nm (spectrum c in Fig.
3B). The last spectroscopic intermediate observed
(C in Reaction 1) had a peak at 412-413 nm (spectrum d in Fig. 3B) and appears similar to the overall
spectrum at the terminal stage of the reaction (spectrum b
in Fig. 3A). Accordingly, the mechanism in Reaction 1 can be
refined as shown in Reaction 2.
|
(Reaction 2)
|
The rates of the two steps in Reaction 2 determined by global fit
were 1.98 and 0.02 s
1, respectively. The inactivation
rate determined for peroxidase activity, 0.4 s
1 (Figs. 1
and 2), was much closer to that for the first spectroscopic transition,
implying that irreversible loss of peroxidase activity accompanies
formation of Intermediate III.

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Fig. 3.
PGHS spectral changes during peroxidase
inactivation. A, kinetic scans obtained from
stopped-flow measurements at 2-nm increments between 390 and 430 nm
during reaction of 1 µM PGHS-1 with 30 µM
EtOOH at 24 °C. SVD-analyzed spectra for 0.5-4 s are shown as
thin lines and those between 10 and 60 s are shown as
dotted lines. The spectra of resting enzyme (a)
and the terminal complex after 3 min (b) are presented for
comparison. Data from the initial 100 ms were excluded from SVD
analysis to minimize the spectral contamination from Intermediate I. Arrows indicate the direction of signal amplitude changes in
the 0.5-4-s reaction period. B, resolved spectra for major
intermediates in the inactivation process. Kinetic scan data from
reaction of 0.67 µM PGHS-1 with 25 µM EtOOH
at 24 °C (0.5-4 s of reaction) were fitted to a three-species,
two-step linear mechanism (Reaction 2) using the global fitting
package, Glint. Spectra are shown for resting enzyme (a),
Intermediate II (b), Intermediate III (c), and
the terminal species (d).
|
|
The spectral changes observed during reaction of PGHS with PPHP and
15-HPETE were very similar to those shown in Fig. 3A for EtOOH (data not shown). As was described for the EtOOH reactions, spectra for two intermediates were resolved by fitting the data to a
two-step process (Reaction 2). The spectrum resolved for the earlier
intermediate (Intermediate III in Reaction 2) had its peak at 415-416
nm for the reactions with PPHP and 15-HPETE, slightly longer than
413-414 nm found for the same intermediate during reaction with EtOOH.
There were no obvious differences observed for the terminal species
with different hydroperoxides.
The double-monochromator Olis rapid scan instrument was used to obtain
spectral data extending to the visible region for inactivation intermediates. Due to the weak absorbance of the intermediates in the
visible region, a higher enzyme concentration (2.5 µM) was employed. Rapid scan spectral data obtained during reaction of PGHS
with EtOOH were consistent with the two-step inactivation process in
Reaction 2 and were fitted to the same mechanism (Fig. 4). The spectral changes observed in the
Soret region with the rapid scan instrument were very similar to those
obtained by kinetic scan at incremental wavelengths using a lower level
of PGHS (0.5 µM), thus confirming that the increased
enzyme concentration did not change the self-inactivation mechanism.
The resolved spectrum for Intermediate III obtained by SVD analysis and
global-fitting of the rapid scan data to Reaction 2 (spectrum
b in Fig. 4) was very similar to that obtained by the kinetic scan
approach (spectrum c in Fig. 3B) both in its
absorption peak position and its intensity. In the visible region,
Intermediate III displayed a single broad peak centered at 560 nm, with
an extinction coefficient of 22.7 mM
1
cm
1. This feature is absent from the spectra of resting
enzyme and the terminal species (Fig. 4), indicating that the heme
electronic configuration in Intermediate III is distinct from those of
other recognized peroxidase intermediates. The rates obtained for the two steps in Reaction 2 from the rapid scan data were 0.46 and 0.032 s
1, respectively. The value for the fast spectral change
here is essentially the same as that for peroxidase inactivation (Fig. 2), supporting the contention that peroxidase inactivation accompanies Intermediate III formation.

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Fig. 4.
Resolved spectrum of Intermediate III in both
Soret and visible regions from rapid scan measurements. PGHS-1
(2.5 µM) in 0.1 M potassium phosphate, pH
7.2, containing 0.1% octyl- -D-glucopyranoside, 0.1%
Tween 20, and 10% glycerol was reacted with 75 µM EtOOH
at 24 °C on an Olis RSM-1000 instrument. Spectra obtained in the
Soret and visible regions were subject to SVD analysis and global
fitting according to the linear model described in Reaction 2. Data for
the first 100 ms were excluded from the analysis to minimize
contamination from Compound I. The resolved spectrum of Intermediate
III (b) is shown together with those of resting PGHS-1
(a) and the terminal species after 8-min reaction
(c), the latter obtained with a Hewlett-Packard model 8453 UV-visible spectrophotometer.
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Analysis of Heme Structural Changes during Peroxidase
Self-inactivation--
Rapid-quench experiments were conducted to look
for covalent changes of the PGHS heme group. For this, PGHS was reacted
with 10 eq of EtOOH, and aliquots were removed at various times for solvent quenching and HPLC analysis. As shown in Fig.
5A, the chromatographic
profiles for the control sample (EtOOH was replaced by buffer) and the
sample obtained at 5 s of reaction were very similar. The main
heme peak, monitored by absorbance at 400 nm, eluted at 20.5 min and
was clearly separated from the PGHS apoprotein peak registered by 220 nm absorbance, which elutes near 26 min. A time-dependent
increase of a 400-nm absorbing species with a 21.7-min retention time
was noticed at expanded scales, but it accounted for only a very small
percentage the total heme. As shown in Fig. 5B, the
integrated area of the main A400 peak changed little with reaction time. Thus, most of the PGHS heme remained chemically intact during the time the peroxidase was inactivated.

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Fig. 5.
HPLC analysis of PGHS heme structural changes
during peroxidase self-inactivation. Equal volumes of PGHS-1 (5.8 µM in 0.1 M potassium phosphate, pH 7.2, containing 0.1% Tween 20, 10% glycerol, and 0.1%
octyl- -D-glucopyranoside) and 60 µM EtOOH
were reacted with at 24 °C for 0, 0.1, 0.2, 0.5, 1.0, 2.0, 5.0, 10, and 60 s and quenched by an aqueous solution of 60% acetonitrile
and 1.2% trifluoroacetic acid. The final concentrations of protein,
peroxide, and the quenching solvents in the quenched sample were 1/3
those of the originals. A, HPLC analysis were performed as
detailed under "Experimental Procedures." As the profiles for every
sample were very similar, only the results from the samples quenched at
0 and 5 s are shown. B, integrated areas for the main
heme chromatographic peak (monitored by A400)
are plotted against reaction time.
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Mathematical Fitting to the Mechanistic Model of Peroxidase
Catalysis in PGHS--
Early kinetic events in the reaction of PGHS
with peroxide, including formation and degradation of Intermediate I,
were not considered in the SVD and global fitting to Reaction 2. In
addition, minor spectral species and reactions involving recycling of
higher oxidation states of heme were ignored to simplify data analysis. To validate these data manipulations, further analysis of PGHS self-inactivation by peroxide was conducted by simulation and fitting
to the more complete mechanism shown in Scheme 1 (Steps 1-8 listed in
Table I) using the SCoP numerical integration program. In this
mechanism, the reaction pathway divides at Intermediate II, with one
route leading to Intermediate III and irreversible inactivation (via
k7) and the other route recycling the enzyme sequentially back to Compound II (or Fe(III) Tyr·) and resting
PGHS (via k3 or k5). The
rate constants for Steps 1 and 2 were taken from published data (4).
The rate constants for the recycling steps, Steps 3-6, were set at
103 to 105 M
1
s
1, in line with literature values for various PGHS
peroxidase cosubstrates (19-22). Five equivalents of endogenous
reductant were assumed in the simulations based on earlier estimates
(25). The simulations indicated that the flux through the recycling
steps is minimal unless exogenous cosubstrate is added. The main
determining factors for the speed of transitions in the 0.5-5-s period
were the values of k7 and
k8. To fit the simulations to the observed
absorbance change kinetics, molar absorbance coefficients for
individual wavelengths were based on the resolved spectra for
Intermediates II and III and terminal species obtained in this study,
and on previously published spectra corresponding to resting PGHS and Intermediate I (4). The floating parameters in the computer fitting
included k7 and k8.
Fittings that floated the k3
to k6 parameters always converged
with rate constant values of less than 105
M
1 s
1. Floating the amount of
endogenous reductant usually converged to values close to 0. These
results indicate a minimal contribution of the reductive recycling
steps (k3 to
k6) to the overall kinetics. Typical kinetic data and
fittings for three different wavelengths are shown in Fig.
6. The initial rapid changes before
0.5 s reflect the formation of Intermediates I and II. The
increase in A392 and decrease in
A428, accompanied by a relatively static
A404, in the 0.1-5-s period coincide with the
conversion of Intermediate II to Intermediate III. The optimal values
for k7 and k8 obtained in
this fitting were 1.87 and 0.01 s
1, respectively. These
values match nicely with those obtained by global fitting to the simple
sequential model represented by Reaction 2, indicating that the fitting
results with the complete mechanism in Table I are essentially the same
as those obtained with the simplified mechanism in Reaction 2.

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Fig. 6.
Computer fitting of single-wavelength kinetic
data to mechanistic model. Kinetic data from the experiment shown
in Fig. 3B are compared with predicted kinetics at the same
wavelengths generated by computer fitting to the mechanism represented
by Steps 1-8 in Table I. The values of rate constants
k1 and k2 were fixed at
1.4 × 107 M 1
s 1 and 320 s 1 (4), respectively;
k3 to k6 were fixed at
105 M 1 s 1, and
endogenous reductant was set to 3 µM (25). Absorbance
coefficients at 404, 392, and 428 nm for each enzyme species were
obtained from the resolved spectra shown in Fig. 3B. The
optimal values obtained for k7 and
k8 were 1.87 and 0.01, respectively.
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 |
DISCUSSION |
The basic kinetics of formation of Intermediate I (Compound I) and
Intermediate II during reaction of PGHS-1 with peroxides are well known
(1-4). Formation of Intermediate I is very rapid, with rate constants
ranging from 108 M
1
s
1 for lipid hydroperoxides to 105
M
1 s
1 for
H2O2 (1-4). Formation of Intermediate II
occurs as a largely unimolecular process, with a rate constant of
50-500 s
1 (3, 4). It is clear from the present results
that both of these processes are much more rapid than the loss of
peroxidase activity, which averaged about 0.4 s
1 (Fig.
2). A second striking aspect of the present results is that
inactivation of the peroxidase is a first-order process, independent of
peroxide concentration and structure. This behavior strongly indicates
that the inactivation reaction step occurs after formation of
Intermediate I or II. Two lines of evidence indicate that the starting
point for inactivation is Intermediate II. First, little Intermediate I
is present during the period when the bulk of self-inactivation occurs
under our experimental conditions. Second, kinetic scan (Fig. 3) and
rapid scan (Fig. 4) data show no evidence of an additional reaction
step before the conversion of Intermediate II to Intermediate III, and
the observed rate constant of this conversion is in the range of that observed for loss of peroxidase activity (Figs. 1 and 2).
Previous examinations of PGHS peroxidase self-inactivation found the
process to be sensitive to peroxide structure and concentration (16,
17), but these studies were done under "steady-state" conditions
with exogenous reductant present. The cosubstrate facilitates recycling
of Intermediate II back to resting enzyme, making the steady-state
level of Intermediate II, and thus the inactivation rate, dependent on
the rate of Intermediate II regeneration by additional peroxide (via
Intermediate I). As a result, the steady-state level of
Intermediate II under such conditions is far from stoichiometric, and
the earlier results are not directly comparable with those from this study.
The proposed inactivation of PGHS peroxidase from Intermediate II
contrasts with the inactivation pathway involving Compound I reported
for the reaction of horseradish peroxidase with excess m-chloroperbenzoic acid (26). Horseradish peroxidase
Compound I is rather stable, whereas PGHS Compound I quickly converts
to Intermediate II. It may well be that this large difference in the
lifetime of Compound I leads to a different inactivation processes in
PGHS and horseradish peroxidase. For horseradish peroxidase, as few as
2 eq of m-chloroperbenzoic acid are sufficient to induce inactivation, permitting calculation of the partition ratio between the
inactivation and recycling processes (26). For PGHS-1, the presence of
endogenous reductant complicates a quantitative analysis because the
endogenous reductant is not titratable by oxidants such as
ferricyanide, and its quantity varies from preparation to
preparation.2 To simplify the
analysis, we did not include exogenous reducing cosubstrate in the
present experiments. This simplified reaction system led to minimal
recycling of the enzyme to resting state via reductive steps. Bakovic
and Dunford (27) observed a rate of 0.08-0.11 s
1 for the
decay of Intermediate II in the reaction of PGHS-1 and m-chloroperbenzoic acid. This rate was zero-order with
respect to the peroxide concentration, a finding very similar to our
results, and their rate constant value falls between the rate constants we obtained for steps 1 and 2 in Reaction 2. The differences between the present results and their studies (27) may partly be due to the
unresolved two reaction steps (k7 and
k8 in Scheme 1) in their data analysis of the
inactivation measurements and the presence of added reductant,
diethyldithiocarbamate, in their reaction mixture.
The present studies have isolated a new spectral intermediate in the
self-inactivation process, i.e. Intermediate III, using both
kinetic scan and rapid scan approaches (Figs. 3 and 4). Intermediate III appears to be produced directly from Intermediate II, which contains Fe(IV)=O and a tyrosyl radical. The spectral changes in the
Soret region during the transition from Intermediate II to III showed
an isosbestic point at 406 nm (Fig. 3A), near that for the
transformation of Intermediate I to Intermediate II (4). However, the
spectral changes in these two transitions are quite distinct, in that
the absorbance changes on the two sides of the isosbestic point
occurred in opposite directions. The overall spectral line shape of
Intermediate III roughly resembles that reported for covalently bound
heme in myoglobin pretreated with hydrogen peroxide (28). However, heme
analysis during the peroxidase inactivation rules out the possibility
of significant covalent linkage between the heme group and the PGHS-1
protein, because no metalloporphyrin chromophore was associated with
protein peak in the HPLC profile (Fig. 5A). Furthermore, the
PGHS-1 heme structure was not changed by peroxidase self-inactivation,
as evidenced by the unchanged chromatographic behavior of the heme
isolated from samples taken at different times during reaction with
EtOOH (Fig. 5). Taken with the observed spectral changes, the
chromatographic results make it likely that the heme group found in
Intermediate III is not properly coordinated to the histidine ligand(s)
but still remains non-covalently associated with the protein.
Irreversible heme destruction can be observed during cyclooxygenase
catalysis (12), but this appears to be a much slower process than
conversion from Intermediate II to III.
Several observations indicate that the process of PGHS peroxidase
self-inactivation begins with Intermediate II. First of all, the rate
of conversion from Intermediate II to Intermediate III, 0.5-2
s
1, is only slightly faster than the decay rate for
peroxidase activity (~0.4 s
1). In fact, measurements
using m-chloroperbenzoic acid instead of
H2O2 in the peroxidase assay gave decay rates
around 1-2 s
1, very similar to the rate for the
Intermediate II to Intermediate III transition (data not
shown).3 The similar rates
for conversion from Intermediate II
Intermediate III measured by
spectral changes and for the decay of peroxidase indicate that they
reflect the same kinetic event. Second, the observed rate for
Intermediate III transformation to the terminal spectral species
(0.01-0.05 s
1) was kinetically quite distinct from the
reaction of Intermediate II
Intermediate III. It is thus concluded
that Intermediate III is itself catalytically inactive.
Reduction of the peroxide-generated radical in crude PGHS by added
cosubstrate was found to decrease the amount of self-inactivation (29).
Various reducing cosubstrates have been reported to extend the
peroxidase activity (15, 19-22), and there was an inverse correlation
between the residual peroxidase activity and the tyrosyl radical
intensity when purified PGHS-1 was reacted with peroxide (18, 30).
These observations indicate the linkage between tyrosyl radical and
self-inactivating damage in PGHS. However, the present analyses of heme
structure (Fig. 5) show clearly that a reaction covalently linking a
tyrosyl radical to the heme moiety is not the cause for peroxidase inactivation.
The relationship of PGHS peroxidase inactivation to cyclooxygenase
inactivation is of considerable interest. Previous measurements of the
cyclooxygenase inactivation rate during reaction with fatty acid (31)
produced a value that is roughly comparable to those obtained for
peroxidase inactivation in this study. Much slower rates for
cyclooxygenase inactivation (1-5 min
1) were reported
earlier by Lands and co-workers (10, 12). Comparison between these
rates for cyclooxygenase and peroxidase inactivation is considerably
complicated by the reaction conditions used for the cyclooxygenase
measurements. The peroxide levels in the cyclooxygenase reactions vary
widely with time and are sensitive to the level of reducing cosubstrate
present (31). In the branched-chain mechanism (3), cyclooxygenase
catalysis requires reaction of the enzyme with peroxide to generate the tyrosyl radical. This predicts that inactivation of the cyclooxygenase by peroxide should be at least as fast as peroxidase inactivation. Further quantitation of the cyclooxygenase self-inactivation will be
needed to test this hypothesis.
 |
ACKNOWLEDGEMENTS |
We thank Drs. John S. Olson and Mark S. Hargrove for their kind assistance in using the Olis RSM-1000 system.
We also thank Dr. Richard J. DeSa at Olis for help in updating and
testing the global fitting program.
 |
FOOTNOTES |
*
This work was supported by U. S. Public Health Service
Grants GM44911 (to A.-L. T.), GM30509 and GM52170 (to R. J. K.), and ES08365 (to Y. O.), and Burroughs Wellcome Fund New Investigator Award
in Toxicology (to Y. O.).The costs of publication of this article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
¶
To whom correspondence should be addressed: Division of
Hematology, University of Texas Health Science Center, P. O. Box
20708, Houston, TX 77225. E-mail:atsai{at}heart.med.uth.tmc.edu.
2
A.-L. Tsai, G. Wu, and R. J. Kulmacz,
unpublished observations.
3
Measurements of the dependence of peroxidase
activity on PGHS-1 concentration (0.01-3.0 µM) revealed
a linear relationship with m-chloroperbenzoic acid as
substrate but a tendency for saturation with
H2O2 as substrate. This indicates that
peroxidase assays with H2O2 slightly
underestimate peroxidase activity, more so when the activity is high
than when it is low. As a result, measurements of surviving peroxidase
activity with H2O2 slightly underestimate the
rate for decay of peroxidase activity.
 |
ABBREVIATIONS |
The abbreviations used are:
PGHS-1, ovine
prostaglandin H synthase-1;
15-HPETE, 15-hydroperoxyeicosatetraenoic
acid;
EtOOH, ethyl hydrogen peroxide;
PGG2, prostaglandin
G2;
PGH2, prostaglandin H2;
PPHP, trans-5-phenyl-4-pentenyl-1-hydroperoxide;
TMPD, N,N,N',N'-tetramethyl-p-phenylenediamine;
HPLC, high pressure liquid chromatography;
SVD, singular value
decomposition.
 |
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