Responses of vascular endothelial oxidant metabolism to lipopolysaccharide and tumor necrosis factor-α

ARCHIVES

OF BIOCHEMISTRY

AND

BIOPHYSICS

Vol. 294, No. 2, May 1, pp. 686-694, 1992

Responses of Vascular Endothelial Oxidant Metabolism to Lipopolysaccharide and Tumor Necrosis Factor-a James A. Royall,* Departments Birmingham,

Paula D. Gwin,*

Dale A. Parks,*+$

of *Pediatrics, TAnesthesiology, $Biochemistry, Birmingham, Alabama 35233-6810

Received July 31,1991, and in revised form January

Press,

Inc.

r To whom correspondence and reprint requests should be addressed at Department of Anesthesiology, University of Alabama at Birmingham, 619 19th St. South, 941THT, Birmingham, AL 35233-6810. 686

University

of Alabama at

8,1992

Quantification of intracellular and extracellular levels and production rates of reactive oxygen species is crucial to understanding their contribution to tissue pathophysiology. We measured basal rates of oxidant production and the activity of xanthine oxidase, proposed to be a key source of 0; and Hz02, in endothelial cells. Then we examined the influence of tumor necrosis factor-a and lipopolysaccharide on endothelial cell oxidant metabolism, in response to the proposal that these inflammatory mediators initiate vascular injury in part by stimulating endothelial xanthine oxidase-mediated production of 0; and HzOz. We determined a basal intracellular HzOz concentration of 32.8 + 10.7 pM in cultured bovine aortic endothelial cells by kinetic analysis of aminotriazolemediated inactivation of endogenous catalase. Catalase activity was 6.72 f 1.61 U/mg cell protein and glutathione peroxidase activity was much lower, 8.13 & 3.79 mU/ mg protein. Only 0.48 * 0.18% of total glucose metabolism occurred via the pentose phosphate pathway. The rate of extracellular HzOz release was 75 + 12 pm01 . min-’ . mg cell protein-‘. Intracellular xanthine dehydrogenase/oxidase activity determined by pterin oxidation was 2.32 + 0.75 pU/mg with 47.1 + 11.7% in the oxidase form. Intracellular purine levels of 1.19 + 1.04 nmol hypoxanthine/mg protein, 0.13 f 0.17 nmol xanthinelmg protein, and undetectable uric acid were consistent with a low activity of xanthine dehydrogenasel oxidase. Exposure of endothelial cells to 1000 U/ml tumor necrosis factor (TNF) or 1 rg/ml lipopolysaccharide (LPS) for 1-12 h did not alter basal endothelial cell oxidant production or xanthine dehydrogenase/oxidase activity. These results do not support a casual role for HzOz in the direct endothelial toxicity of TNF and LPS. o 1992 Academic

and Bruce A. Freeman**t’$$l

and $Physiology and Biophysics,

Endothelial cells have been proposed to be both significant sources of oxidants (l-3) and targets of oxidant injury (4, 5) in inflammatory diseases. Toxic bacterial products, such as LPS,’ can directly injure endothelial cells (6) with recent evidence suggesting that cytokines, particularly TNF released by LPS-stimulated mononuclear phagocytes, are important mediators of the in vivo effects of LPS (7). Exposure to LPS and TNF directly impairs the barrier function of endothelial monolayers (8), and LPS ultimately induces morphologic changes and cell death (6). However, endothelial cells are more than passive targets in acute microvascular injury associated with inflammation. Under the influence of inflammatory mediators, endothelial cells express cell surface receptors which increase neutrophil adherence (9) and are more susceptible to neutrophil-mediated cytotoxicity (10). Endothelial cells are also proposed to be a source of iron for hydroxyl radical formation from neutrophil-derived 0, and Hz02 via the Fenton reaction (11). Finally, endothelial cells can participate alone or in concert with reticuloendothelial cells in the generation of . NO (endothelial-derived relaxation factor) and O,, yielding the potent oxidant ONOO- (12,13). Endothelial cell reactive oxygen species production and metabolism are proposed as important factors in acute vascular injury in several pathologic states and may also play a regulatory role in some cell’ functions (14). Previous reports have emphasized the cytotoxic effect of Hz02 toward endothelium in acute inflammation (4, 5, 11). Whether enhanced endogenous endothelial production of

2 Abbreviations used: LPS, lipopolysaccharide; TNF, tumor necrosis factor-a; AT, 3-amino-1,2,4-triazole; (aminotriazole); XDH, xanthine dehydrogenase; X0, xanthine oxidase; GSH, glutathione; GSSG, oxidized glutathione; HEPES, N-2-hydroxyethylpiperazine-Nr-2-ethanesulfonic acid; MES, 2-[N-morpholinolethanesulfonic acid, PMSF, phenylmethylsulfonyl fluoride; DTT, dithiothreitol; DTPA, diethylenetriaminepentaacetic acid; CHAPS, 3-[(3-cholamidopropyihlimethylammonio]propanesulfonic acid; DMEM, Dulbecco’s modified Eagle’s medium. 0003.g&31/92 $3.00 Copyright 0 1992 by Academic Press, Inc. All rights of reproduction in any form reserved.

INFLAMMATORY

MEDIATORS

AND

ENDOTHELIAL

reactive oxygen species has a causative role in direct inflammatory mediator-induced injury is unclear. This mechanism of injury is indirectly supported by several previous reports. Conversion of XDH to the oxygen radical producing form, X0, occurs in rat pulmonary artery endothelial cells exposed to activated neutrophils (2), 5a, TNF, and N-formyl-Met-Leu-Phe (3). Enhanced extracellular release of 0; has been reported from human umbilical vein endothelial cells exposed to interleukin-1 (15) and enhanced release of 0; and H202 has been reported during endothelial phagocytosis (16). Increased endogenous production of reactive oxygen species has been suggested as a mechanism for LPS-induced injury in bovine pulmonary artery endothelial cells (1). This conclusion was based on a relative decrease in reduction of nitro blue tetrazolium and decreased oxidation of 2’,7’-dichlorofluorescein diacetate, both nonspecific markers of oxidant production, in dimethylsulfoxideand allopurinol-treated cells which had been previously exposed to LPS. However, direct quantification of changes in cellular reactive oxygen species production in TNF- or LPS-treated cells has not been reported. Sensitive and unambiguous techniques for quantification of intracellular levels or rates of production of reactive oxygen species are crucial for developing a mechanistic understanding of the physiologic and pathologic roles of reactive oxygen species. However, few such methods have been described due to confounding factors, including competing reactions between oxygen radicals and endogenous substances, such as antioxidants, and oxygen radical-independent oxidation/reduction reactions of intracellular probes. In this study, we assessed a portion of endogenous intracellular HzOz production and metabolism, which indicates spontaneous and enzymatically catalyzed 0, dismutation as well as the divalent reduction of molecular oxygen, under basal conditions and after exposure to TNF or LPS. Intracellular H202 can also diffuse extracellularly or be scavenged by glutathione peroxidase, yielding GSSG which is rereduced by NADPH and glutathione reductase (17). We also assessed the rate of extracellular release of H202 as well as the activity of glucose utilization by the pentose phosphate shunt, shown to be an indirect measure of enhanced rates of HzOz reduction by NADPH-dependent processes (18). Since X0-derived reactive oxygen species have been reported in vascular endothelium (19), an estimation of the contribution of X0 to net cell Hz02 production and the influence of LPS and TNF was evaluated. EXPERIMENTAL

PROCEDURES

Materials. Methylene blue, 2-amino-4-hydroxy pteridine (pterin), isoxanthopterin, horseradish peroxidase, p-hydroxyphenylacetic acid, allopurinol, aminotriazole, HEPES, Hoechst 33258, MES, PMSF, DTT, calf thymus DNA, methylbenzethonium hydroxide, menadione, ammonium molybdate, DTPA, glutathione reductase, t-butyl hydroperoxide, reduced glutathione, and NADPH were obtained from Sigma Chemicals, St. Louis, Missouri. Leupetin was from Boehringer-Mann-

OXIDANT

METABOLISM

687

heim Biochemicals, Indianapolis, Indiana. Bovine liver catalase was from Worthington Biochemical Corp., Freehold, New Jersey, and CHAPS from Calbiochem Co., La Jolla, California. Radiolabeled glucose ([l“C]glucose and [6-i4C]glucose) was from Amersham Corp., Arlington Heights, Illinois. Culture plasticware was from Costar, Cambridge, Massachusetts. Wells for CO, traps were from Kontos Corp., Vineland, New Jersey. Medium 199, trypsin-EDTA, antibiotic-antimycotic solution, and phenol red-free Medium 199 were obtained from GIBCO Laboratories, Chargin Falls, Ohio. Fetal calf serum was obtained from Hyclone Laboratories, Logan, Utah. Ryan’s growth supplement was obtained from Dr. U. Ryan, Monsanto Inc., St. Louis, Missouri. Equipment used included a cell sonifier (Sonifier Cell Disruptor, Heat SystemsUltrasonics, Plainview, NY), spectrophotometer (Gilford Response UVVis spectrophotometer, Ciba-Corning Diagnostic Corp., Medtield, MA), spectrofluorometer used in DNA determinations (Farrand System 3 scanning Auorometer, Optical Technology Devices, Elmsford, NY), spectrofluorometer used in xanthine dehydrogenase and oxidase determinations (SLM SOOOC,SLM Instruments Inc., Urbana, IL), and liquid scintillation counter (LKB-Wallac 1214 Rackbeta, Finland). Highpressure liquid chromatography equipment (autosampler, UV diode array detector, and Omega data acquisition system) were from Perkin-Elmer Corp., Norwalk, Connecticut. Human recombinant tumor necrosis factor-a (6.24 X 10s U/ml specific activity as determined by mouse L-929 fibroblast lysis assay, with LPS contamination being 3.2 X 10m6ng/lOOO U by Limulus amoebocyte lysate assay) was supplied by Dr. L. Lin, Cetus, Sunnyvale, California. Repeat fibroblast lysis assay performed after completion of these studies showed no loss of TNF activity. Lipopolysaccharide W (Escherichia Coli 055: B5) was from Difco Laboratories, Detroit, Michigan. Cell culture. Bovine aorta acquisition, endothelial cell isolation, and cell culture were performed as previously described (8). Endothelial cells were maintained in Medium 199 with 1% antibiotic-antimycotic solution, 5% fetal calf serum, 5% Ryan’s growth supplement, and 2.5 rig/liter ammonium molybdate. Fetal calf serum and Ryan’s growth supplement can serve as oxidant scavengers (20) and therefore were excluded from the medium during biochemical analyses. Studies were performed on cell monolayers 1 to 3 days after reaching confluence. Cells were used up to the 15th passage. In each study, controls were performed concurrently with experimental conditions and all cells were split from the same parent flask. Aminotriazole-mediated inactivation of endogenous cataluse actiuity. Intracellular H202 concentration was determined by analysis of the rate of AT-mediated inactivation of endogenous catalase activity (21). Catalase inactivation studies were performed using cells grown in 25-cm2 culture flasks, with cells from two flasks combined for each time point. Endothelial cell monolayers were incubated with 10 mM AT in phenol red-free Medium 199 and 10 mM Hepes, pH 7.4, at 37°C with the exception of the t = 0 flasks. After the desired incubation period in the AT solution, monolayers were washed, 1 ml of 50 mM potassium phosphate, 0.1 mM EDTA, and 0.1% CHAPS, pH 7.0, was added to each flask, and the cells were removed by scraping. Cells from the two 25-cm2 flasks were placed in a single tube, lysed by sonication, and immediately placed on ice. Cell lysate catalase activity was measured spectrophotometrically by the method of Bergmeyer (22) based on the disappearance of 10 mM H,Op at 240 nm (extinction coefficient of 43.6 Mm’ cm-i). One unit of catalase activity was defined as that decomposing 1 pmol H,O,/min at 25°C. Cell protein was determined by the method of Lowry et al. (23) with an albumin standard. Cell DNA was determined by fluorometric assay as described by Labarca et al. (24) with calf thymus DNA as the standard. For most studies, cell lysate catalase activity was determined after 8 min AT incubation intervals over a IO-min period. In some studies, incubation with AT was extended to 4-6 h to determine the kinetics and extent of inactivation of catalase. The steady-state H202 concentration was calculated in a similar fashion to that described in studies of rat brain (25). Figure 1 shows catalase/H,Oz metabolism. When AT concentration is sufficiently high and not rate limiting, the steady-state concentration of H,On can be calculated from

688

ROYALL

[H,O,]

= F

1

where kti is the pseudo-first-order rate constant of catalase inactivation determined from the experimentally derived AT-mediated catalase inactivation curve and the value of kl is known (1.7 X 10’ M-la-‘) (see Miniprint Supplement). In two studies, endothelial cells were exposed to 95% O2 plus 5% COz or to exogenously added 100 pM HzO, for 1 h prior to determination of kmt. Basal H202 concentrations were 12.3 and 10.2 pM, with hyperoxia were 19.1 and 20.8 pM, and with exogenous H202 were 37.6 and 43.4 pM (a 55 and 104% increase with hyperoxia; a 206 and 325% increase with exogenously added H,O,). release of H202 was Extracellular Hz02 release. The extracellular determined by fluorometric assay (26). In 25-cm’ flasks, 5 ml of medium consisting of phenol red-free Medium 199 with 1.9 g/liter sodium bicarbonate, 1 mM Hepes, 1.6 mM p-hydroxyphenylacetic acid, and 95 rg/ ml horseradish peroxidase, pH 7.4, was added. To quantify the rate of H,Os release into the extracellular compartment, sequential measurements of HzO,-mediated oxidation ofp-hydroxyphenylacetic acid in cell culture medium were made over a 2-h period. An 18 to 24% decrease in extracellular Hz02 detection was noted in the presence of 300 U/ml bovine liver catalase, consistent with expected partial catalase competition with horseradish peroxidase for H202. Exposure to 1 nM menadione for 15 min increased extracellular Hz02 release from a basal rate of 31.2 + 2.6 to 475 + 53.8 pmol * min-’ * mg cell protein-’ (X + SD, n = 4). Differential production of raPentose phosphate pathway activity. diolabeled CO2 from [1-‘“Clglucose and [6-‘4C]glucose was measured (27). Approximately 0.68 &i (12 pmol glucose) of either [1-“Clglucose or [6-“Clglucose (specific activities of 55.8 mCi/mmol) in 4 ml of medium (5.6 mM glucose) was added to 25-cm* culture flasks. Immediately, 100 pl of medium was removed for determination of glucose specific radioactivity. A COP trap consisting of a plastic well containing methylbenzethonium hydroxide and a 25 X lo-mm filter paper wick was placed in the flask and then sealed. After 6 h, 150 ~1 of medium was removed and one drop of 20% trichloroacetic acid added to purge dissolved COz before 100 ~1 of medium was counted. The difference between the initial medium sample and the 6-h sample represented net glucose metabolism. An addition of 300 ~1 of 20% trichloroacetic acid was made to the remaining medium in flasks and 14C02 was trapped for 16 h. The specific yield of CO1 from [1-“Clglucose (Glco,) and from [6-‘“Clglucose (GGco,) was determined from the CPM of trapped COz divided by the decrease in CPM of medium over the 6-h study period. If glucose was metabolized only via the Embden-Meyerhof pathway and the pentose phosphate pathway (assumes no metabolism by nontriose-P pathways such as glycogen) the portion of total glucose metabolism occurring via the pentose phosphate pathway (PC) is calculated from

Glcoz - G~co,--_ 1 - GGco,

ET AL. and lysed by sonication. The lysates were immediately frozen at -86°C in isopentane. Cell lysate XDH + X0 activity was determined at 37°C using a Auorometric assay (345 nm excitation, 390 nm emission) based on pterin oxidation with methylene blue as the electron receptor for combined oxidase and dehydrogenase activity (30). Allopurinol(l0 PM) was added to document specificity of the assay for XDH + X0 and 0.1 ELMisoxanthopterin added as an internal fluorescence standard. One unit of activity is equivalent to oxidation of 1 hmol pterin/min. Intracellular purine concentrations. To measure intracellular hypoxanthine, xanthine, and uric acid levels, cells were deproteinized by addition of cold 11 M perchloric acid (final concentration, 0.44 M) and kept at room temperature for 10 min. Samples were centrifuged at SOOOg for 10 min and the supernatant was frozen at -20°C. Prior to chromatography, samples were adjusted to pH 4.5 by addition of 2.5 M KOH and 0.05 M MES. Standard solutions of xanthine, hypoxanthine, and uric acid were prepared in 30 mM potassium phosphate at pH 4.5. Separations were performed isocratically on a Perkin-Elmer 15 X 0.46-cm column packed with C-18 pecosil (particle diameter, 5 rrn) and a guard column. The mobile phase was 30 mM potassium phosphate buffer containing 0.2% methanol. Xanthine, hypoxanthine, and uric acid were identified by comparison with standards. Diode array detection confirmed purine peaks by characteristic absorption spectra. Concentrations were assessed using internal standards. Results are presented as the mean + SD. StaStatistical analysis. tistical analysis was performed using two statistical software packages (Statgraphics 3.0, Rockville, MD, and Primer, McGraw-Hill, NY). Twoway analysis of variance with the Tukey post hoc test, one-way analysis of variance with the Bonferroni post hoc test, and a two-tailed t test were applied to appropriate studies. A P < 0.05 was considered to indicate a statistically significant difference. Enzyme kinetic analysis was performed with Enzfitter 1.05, Elsevier-Biosoft, Cambridge, UK.

RESULTS Intracellular Hz02 levels The rate of AT-mediated inactivation of endogenous endothelial cell catalase activity

CATALASE

t %02

I

kl

COMPOUND

I

3PC 1+ 2PC ’

Exposure to 95% O2 plus 5% COP or to 50 pM t-butyl hydroperoxide for 1 h increased PC from 0.22 f 0.17% under basal conditions to 1.58 + 0.11% with hyperoxia and 9.38 + 1.77% with t-butyl hydroperoxide (X rt SD, n = 4). Enzyme analysis. Cell glutathione peroxidase activity was determined spectrophotometrically (28) with 5 PM Hz02 as substrate. In some studies, glutathione peroxidase activity using 70 WM t-butyl hydroperoxide as substrate was also determined. The K,,, of glutathione peroxidase for H202 is 1 pM (29); thus enzymatic rates determined with 5 pM Hz02 are 83% of maximal according to Michaelis-Menten kinetics and measured activity was adjusted to reflect V,, glutathione peroxidase activity. For measurement of XDH + X0 activity, cells were removed from flasks by scraping and centrifuged, and pellets were resuspended in cell lysing buffer plus 1 mM PMSF, 0.5 pg/ml leupetin, and 10 mM DTT,

INACTIVE CATALASE

CATALASE 1 CATALASE +

t CH&HO

t 2H20

w3+02

FIG. 1. Catalase/H,O,/aminotriazole interactions. Cat&se typically reacts with H202 to form Compound I. Compound I may (a) react with AT which irreversibly inactivates catalase, (b) react catalatically with H202, or (c) react peroxidatically with an alcohol (ethanol). The rate of Compound I formation (k,) is 1.7 X lo7 M-’ s-l, kAT is the rate of Compound I degradation in the presence of AT, & is the rate of degradation in the presence of Hz02, and ketoh the rate of degradation in the presence of ethanol (reproduced with modifications with permission from Ref. (25)).

INFLAMMATORY 0.12

T

O-O 0-O

0.10 -9 n

0.08~-

7 2 ;

0.06~-

MEDIATORS

AND

ENDOTHELIAL

Control TNF

0.04-0.02 -0.004 -5

5 Aminotriazole

15 25 Incubation Time

35 (min)

I 45

inactivation of intracellular catalase FIG. 2. Aminotriazole-mediated activity. Intracellular catalase activity of endothelial cells (U CAT/ag DNA) was determined over time in the presence of 10 mM AT. Values at each time point represents Z + SD for triplicate measurements. The pseudo-first-order rate constant (k,,) was determined for each curve and intracellular H,O, concentration calculated. In this study, cells were exposed to 1000 U/ml TNF for 4 h. Control intracellular Hz02 concentration was 38.3 pM and TNF-exposed intracellular HzOz concentration was 31.7 pM.

followed single exponential decay kinetics (Fig. 2). Virtually all endogenous catalase activity was inactivated by AT, with only 12.5 + 1.0 mU/pg DNA (0.52 * 0.44 mU/ mg protein) catalase activity remaining after 6 h of incubation with AT (n = 4). The assumption that the AT concentration is sufficiently high such that it is not rate limiting and does not change during the study period is supported by the fit of the experimental data to a single exponential decay model. The validity of our mathematical approach was verified using a cell-free model system which mimicked the cell lysate conditions with Hz02 and catalase concentrations similar to those measured in cell studies. Exposure times and TNF concentrations were based on our previous work with bovine aortic endothelial cells, where a TNF-induced increase in endothelial monolayer permeability occurred without gross morphologic changes or increased release of intracellular LDH or [14C]adenine prelabel (8). In these previous studies, we found the maximum TNF-induced increase in permeability to occur after 12 h of exposure to 1000 U/ml TNF. Therefore, exposure times up to 12 h were evaluated. For LPS studies, significant morphologic alterations and cell death were noted after 12 h of exposure to 1 pg/ml LPS, therefore an additional time point at which minor morphologic alterations were first apparent (6-8 h) was evaluated for oxidant production. Cell lysate protein levels (0.32 + 0.05 mg/ml), DNA levels (13.18 + 2.65 pg/ml), and protein/DNA ratios (pg/wg, 23.21 f 5.92) were not altered by experimental conditions or exposure to TNF or LPS. Ceil lysate catalase activity was indexed to both protein and DNA concentration, yielding similar estimations of H202 concentra-

OXIDANT

689

METABOLISM

tion. Under control conditions, the mean intracellular HzOe concentration for combined control exposure groups was 32.8 * 10.7 pM with exposure to TNF or LPS having no significant effect (Table I). Another possible effect of enhanced intracellular rates of reactive oxygen species production could be a decrease in intracellular catalase activity due to direct inactivation by 0, (31). This did not occur in these studies, because control catalase activity (5.72 -t 1.61 U/mg protein) was not affected by exposure of cells to TNF or LPS (data not shown). Pentose phosphate pathway activity/glutathione peroxidase activity. The glutathione peroxidase-reductase system is a major intracellular scavenger of HzOz in some cells (17). During glutathione peroxidase-mediated reduction of H202, GSSG formation requires glutathione reductase-mediated regeneration of GSH using NADPH supplied primarily by the pentose phosphate pathway. The effect of 1000 U/ml TNF or 1 pg/ml LPS on the proportion of total glucose metabolism occurring via the pentose phosphate pathway (PC) was evaluated in two separate studies. The combined results (3c t SD) were control = 0.48 f 0.18% (n = lo), TNF = 0.40 + 0.23% (n = 12), and LPS = 0.35 + 0.16% (n = 12). There was no difference in PC values over the 6-h exposure to TNF or LPS. Intracellular glutathione peroxidase activity was 8.13 t 3.79 mU/mg cell protein (n = 4) with H202 as substrate and 34.6 t- 5.0 (n = 2) with t-butyl hydroperoxide as substrate. Extracellular H202 release. In initial studies, controls were separate flasks evaluated simultaneously with flasks exposed to 1000 U/ml TNF or 1 pg/ml LPS. In three TABLE

Intracellular

I

H,Oz Concentrations and the Influence of LPS and TNF

Exposure time (h)

[H,O,l Control

1 4* 12

38.7 t 11.2 37.8 i 13.7 30.2 i 3.7 Control

1 4 6-8 12

20.0 37.7 31.2 32.6

f + IL +

5.2 8.8 3.7 15.5

(PM) TNF 36.8 rt 14.2 39.8 -c 12.7 29.8 + 13.0 LPS 13.8 34.0 22.8 32.2

f + f -+

6.3 8.8 8.8 7.7

Note. Intracellular H,O, concentration was determined from the rate of aminotriazole-mediated inactivation of endogenous catalase activity. Bovine aortic endothelial cells were maintained in Medium 199 with 1000 U/ml tumor necrosis factor (TNF) or 1 rig/ml endotoxin (LPS). Values are .%i SD with n = 3 except that indicated with an * where n = 4. Statistical analysis was by two-way ANOVA with the Tukey post hoc test.

690

ROYALL

ET AL.

TABLE II Extracellular Endothelial

TABLE III

Cells with

Sequential

Evaluation

Control TNF LPS

Baseline period

70 k 12 66 + 14 77 + 25

Postaddition period 97 2 11 100 f 18 110 & 20

Oxidase Activities

of Controls

H202 release (pmol * min-’ * mg protein-‘) Experimental conditions

Endothelial Cell Xanthine Dehydrogenase and

H202 Release by LPS- and TNF-Treated

Difference (postaddition - baseline)

27+12 34+ 9 33 I!z15

Note. In these studies each flask served as its own control. After a baseline measurement period of 80 min, an addition of 1000 U/ml TNF, 1 pg/ml LPS, or buffer alone (control) was made and analysis continued for an additional 80 min. Values are X + SD, n = 6, and are a summary of two different studies. Under all experimental conditions, postaddition values are different from baseline and therefore comparison of the difference between measurement periods was made by one-way ANOVA.

studies, TNF or LPS was added at the beginning of the 2-h analysis period. The combined results in pm01 0min-’ * mg cell protein-l (5 f SD) were control = 84 f 11 (n = 6), TNF = 89 + 10 (n = 9), and LPS = 79 + 17 (n = 9). In another group of three studies, endothelial cells were exposed to TNF or LPS for 4 h preceding and during the 2-h analysis period. The combined results were control = 72 + 11 (n = 9), TNF = 70 f 19 (n = 9), and LPS = 80 f 15 (n = 7). Exposure to TNF for LPS had no,effect on extracellular H202 release in these studies. To assure that flask-to-flask variation in basal HzOz release would not mask any modest effect of TNF or LPS exposure on extracellular Hz02 release, sequential studies were performed in which each flask served as its own control (Table II). These results are a summary of two separate experiments. After baseline measurement of H202 release for 80 min, TNF, LPS, or control buffer was added. Evaluation of all values (baseline period and postaddition period) for a given experimental condition (control, TNF, or LPS) by least-squares regression analysis yielded correlation coefficients > 0.99. However, rates of extracellular Hz02 release were uniformly higher for both control and exposed monolayers during the postaddition period compared to the baseline period. Therefore, the difference between the rate of H,O, release after mediator addition and that at baseline was the most appropriate comparison. From this, there was no effect of TNF or LPS on extracellular Hz02 release. The mean value for extracellular HzOz release under control conditions was 75 f 12 pm01 . min-l . mg protein-l. Xanthine ox&se, xanthine dehydrogenase, and intracellular purines. We evaluated endothelial cell specific X0 activity, the proportion of total XDH + X0 activity due to X0, and the intracellular level of the substrates hypoxanthine and xanthine and the product uric acid.

Activity Exposure time (min)

Experimental conditions

30

Control TNF LPS Control TNF LPS Control TNF LPS

60 120

x0

XDH

0.68 310.05 1.23 + 0.67 1.00 + 0.69

1.63 f 0.93 1.10 1.25 1.00 1.63 1.05

f + + f +

(hU/mg

0.45 0.31 0.22 0.49 0.42

protein)

+ X0

2.03 + 0.31 2.48 + 0.77 2.18 f 0.84

2.83 f 1.04 2.15 + 0.59

2.30 + 0.20 2.10 + 0.63 2.98 -+ 0.63 1.88 f 0.67

%X0

36 + 6.8 48 f 13.2 44 + 13.8 55 zk 9.8 53 + 12.3 54 + 12.4 50 f 9.6 54 + 4.9 55 + 4.3

Note. Exposure times to 1000 U/ml TNF or 1 pg/ml LPS are indicated Values represent X + SD, n = 4. One unit of activity is defined as the oxidation of 1 pmol pterin/min. Statistical analysis by two-way ANOVA found no significant effect of time or exposure.

The control XDH + X0 activity for all studies was 2.32 f 0.75 pU/mg cell protein, with 47.1 f 11.7% of activity in the oxidase form (Table III). Exposure to TNF or LPS had no effect on net XDH + X0 activity or percentage of total activity due to X0. Intracellular purine levels are given in Table IV. No significant differences in purine levels were noted with TNF or LPS exposure. DISCUSSION We evaluated endogenous H202 production and metabolism in cultured bovine aortic endothelial cells under basal conditions and after exposure to TNF or LPS. From the rate of AT-mediated inactivation of endogenous catalase activity, we determined the concentration of intracellular HzOz to be approximately 32.8 pM under control conditions. Intracellular H202 concentration was not al-

TABLE IV Endothelial

Cell Purine Concentrations Purine concentration

Experimental conditions Control TNF LPS

Hypoxanthine 1.19 f 1.04 2.12 f 0.18

2.28 f 0.67

(nmol/mg)

Xanthine 0.13 + 0.17

0.08 + 0.16 0.08 f 0.16

Uric acid ND ND ND

Note. ND, nondetectable. Cell lysate concentrations of xanthine, hypoxanthine, and uric acid were determined by HPLC. Endothelial cells were exposed to 1000 U/ml TNF or 1 pg/ml LPS for 6 h. Units are nmol/mg cell protein. Values represent 5 f SD, n = 4. Statistical comparison of each purine concentration under different experimental conditions by one-way ANOVA showed no statistically significant differences.

INFLAMMATORY

MEDIATORS

AND

ENDOTHELIAL

tered by exposure to TNF or LPS. Our estimation of intracellular HzOz concentration is less than previous estimations of lo-’ to 10d7 M Hz02 in hepatocytes (32) and greater than the basal HzOz concentration of 3.85 pM calculated for rat brain using an approach similar to that used in our studies (25). Direct comparison of in vitro results to whole organ measurements may not be appropriate, if differentiated cell characteristics change in u&o. However, one explanation for higher endothelial cell Hz02 levels compared to those in whole brain could be heterogeneity of endogenous reactive oxygen species production rates among different cell types. In cultured porcine aortic endothelial cells, approximately 19% of total oxygen consumption was noted to be cyanide-resistant, while less than 4% of total oxygen consumption was cyanide-resistant in lung slices (33). This suggests that endothelial cells may have a greater rate of nonrespiratory oxygen consumption, which may include greater basal rates of oxygen radical production, than other cell types (34, 35). Aminotriazole-mediated catalase inactivation measures the steady-state H202 concentration in the region of the peroxisome. A concomitant knowledge of alternative routes of H202 metabolism in cells, particularly the activity of the glutathione peroxidase-reductase system, extracellular diffusion of HzOz, and intracellular reactions of HzOz, is necessary to interpret results obtained from AT-mediated catalase inactivation (21). The balance between intracellular HzOz production and consumption by scavenging and target molecule reactions will determine if a gradient for H202 diffusion is created such that the concentration of HzOz becomes elevated in other cellular compartments (36). The glutathione peroxidase-reductase system is a key cytosolic antioxidant mechanism. If increased H202 production were within the range that could be metabolized by glutathione peroxidase, the steady-state Hz02 concentration in the region of the peroxisome may be unaltered and thus would not be detected by the AT method. Organ and cell differences exist in the relative contributions of catalase and glutathione peroxidase to Hz02 metabolism (17). We measured the activity of glutathione peroxidase and glucose flux through the pentose phosphate pathway as an indication of HzOz metabolism by this pathway (27). Oxidative stress, NADPH depletion, and elevation in tissue GSSG will cause increased glucose flux via glucose-6-phosphate dehydrogenase in a variety of tissues and cell types (37,38). In support of this concept, stimulation of pentose phosphate pathway activity has been specifically linked to HZOP metabolism by glutathione peroxidase in bovine microvascular endothelial cells (18). There are no previous reports regarding the percentage of glucose metabolism occurring via the pentose phosphate pathway in endothelial cells. In rat lung, 11% of glucose metabolism occurred via the pentose phosphate pathway (39) and in studies of rat adipose tissue this ranged from 7 to 25% (40). Under basal conditions, we determined

OXIDANT

METABOLISM

691

that a low percentage, 0.48% of total glucose metabolism, occurred via the pentose phosphate pathway in bovine aortic endothelial cells. There was no increase in glucose metabolism via the pentose phosphate pathway following LPS or TNF exposure. This suggests a low basal Hz02 production rate or a low specific activity of enzymes involved in this pathway of peroxide metabolism, including glucose-6-phosphate dehydrogenase, 6-phosphogluconate dehydrogenase, glutathione reductase, and glutathione peroxidase. Heterogeneity in intracellular peroxidase systems and their role in protection against H202 in cultured endothelial cells from different sources is well documented (5,41). Additionally, culture conditions such as the availability of selenium, necessary for synthesis of selenium-dependent glutathione peroxidase, and passage number are possible variables between otherwise similar cells (20). Selenomethionine supplementation of porcine aortic endothelial cells increased glutathione peroxidase activity and modulated HzOZ-mediated cell toxicity (42). Serum-supplemented Medium 199 culture medium has less than 2 rig/ml selenium, well below human blood selenium levels of 55 to 143 rig/ml (43). Glutathione peroxidase activity in bovine aortic endothelial cells used herein was 8.13 + 3.79 mU/mg protein when using H202 as substrate and 34.6 +- 5.0 mU/mg protein for t-butyl hydroperoxide, similar to previous reports of glutathione peroxidase activity in endothelial cells isolated from bovine pulmonary arteries (44,45), porcine aorta (20), and porcine pulmonary artery (46) but significantly less than other studies of endothelial cells from porcine aorta (42), human umbilical vein (43), and rat liver (47). The greater glutathione peroxidase activity measured with t-butyl hydroperoxide versus H202 as substrate is consistent with a nonselenium-dependent glutathione peroxidase specific toward organic hydroperoxides but not H202 (48). The relatively low activity of the endothelial cell pentose phosphate shunt and glutathione peroxidase suggests that cataiase may be more important for HzOZ scavenging in bovine aortic endothelial cells. Neither intracellular HZOZ concentration determined by AT inactivation of catalase nor the activity of the pentose phosphate pathway was altered by exposure to TNF or LPS. Therefore, TNF and LPS did not induce an increased rate of Hz02 metabolism by either catalase or the glutathione peroxidase-reductase system in bovine aortic endothelial cells, indicating that cytosolic and peroxisomal HzOz concentration was not affected by exposure to these inflammatory mediators. A significant portion of endothelial cell H202 production was detected extracellularly, thus not reacting with intracellular target molecules and escaping intracellular antioxidant defenses. We hypothesized that increased rates of intracellular H202 production may be reflected in enhanced rates of extracellular H202 diffusion. We observed LPS and TNF to have no effect on the endogenous rate of extracellular release of HzOz. From these results, it can also be questioned

692

ROYALL

whether LPS- and TNF-derived oxidants will stimulate formation of lipid hydroperoxides, previously inferred from observation of conjugated diene release from LPStreated bovine endothelium (1). We have recently observed that conjugated diene formation is a very insensitive marker of excess 0, and HzOz production and can inadvertently reflect enzymatic metabolism of arachidonic acid to vasoactive mediators or cellular release of other chromophores interfering with the UV absorbance indicative of conjugated dienes (13). Recent publications (l-3) suggested that X0 may be a source of increased endogenous reactive oxygen species production in endothelial cells exposed to inflammatory mediators. Thus, we evaluated this enzyme system in the context of our assessment of rates of endothelial Hz02 production after exposure to LPS and TNF. We observed that X0 specific activity and the relative percentages of XDH and X0 activity were unaltered by TNF or LPS exposure. If intracellular concentrations of purine substrates for X0 were to increase secondary to inflammatory mediator exposure, there could be increased endothelial 0; and Hz02 derived from X0, since endogenous purine concentrations are less than the Km of X0 (49). We measured intracellular levels of xanthine, hypoxanthine, and uric acid and found no change in their concentration after TNF or LPS exposure. Our results are in accord with a recent report of intracellular X0 in bovine pulmonary artery endothelial cells (50). These investigators also found no change in absolute X0 activity or the percentage of XDH + X0 activity due to X0 after exposure to 10 pg/ml LPS for up to 8 h. Although LPS-mediated endothelial cell toxicity was reduced by deferoxamine, this was not due to iron chelation causing a reduction in hydroxyl radical formation. This highlights the difficulties in drawing conclusions concerning mechanisms of oxidant injury from inhibitor studies alone as opposed to direct measurement of reactive oxygen species and analysis of reaction by-products. Our results are not in agreement with a recent report that 10 nM TNF induced the conversion of XDH to X0 in rat pulmonary artery microvascular endothelial cells (3). Methodological differences may be a factor in the discrepant results. In our studies, the TNF exposure was somewhat lower (1000 U/ml = 2.45 nM). We excluded serum from our study medium to decrease the oxygen radical scavenging effect of the medium itself (20) while the study of rat pulmonary artery microvascular endothelial cells was performed in the presence of serum. Fetal calf serum has been documented to enhance TNF- and LPS-induced increased endothelial monolayer permeability (6,8) and it is possible that serum is needed for a significant inflammatory mediator effect on XDH + X0. Another factor may be interspecies heterogeneity of XDH + X0 activity and its variable potential for a pathologic role among different types of endothelial cells maintained under different culture conditions. The activity of XDH

ET AL.

+ X0 in the study of rat pulmonary artery microvascular endothelial cells was expressed as nmol uric acid formed/ lo6 cells/min and cannot be compared directly with our XDH + X0 activity assay, which is based on pterin oxidation. Pterin is oxidized more slowly than xanthine depending on temperature and source of the enzyme, with ratios of xanthine to pterin oxidation rates ranging from 2.2 to 8.5 (30). Assuming the highest xanthine/pterin oxidation ratio and expressing both results as pmol xanthine oxidized/min/75 cm2 cell surface area (U/75 cm2), the XDH + X0 activity of our bovine aortic endothelial cells was 19.7 pU/75 cm2 and for rat pulmonary artery microvascular endothelial cells was 46.8 mu/75 cm2. Even considering inaccuracy due to assumptions made for this comparison, it is clear that the XDH + X0 activity of rat pulmonary artery microvascular endothelial cells is substantially greater than bovine aortic endothelial cells and is less reflective of human tissues, which also have relatively lower X0 specific activities (30, 49). Another potential factor in the variation of XDH + X0 activity is vessel source. Immunolocalization studies suggest that large vessel endothelium does not contain significant X0 activity while X0 activity is detectable in capillary endothelium (51, 52). However, this finding is in conflict with other information noting detectable X0 activity in large vessel endothelium by histochemical techniques (51). Culture conditions can also affect cellular XDH + X0 activity. Bovine aortic endothelial cells maintained in a DMEM-based culture medium have a sixfold higher XDH + X0 activity than those maintained in a Medium 199based culture medium (53). A typical control cell X0 activity of 1.1 f 0.64 pU/mg protein, if saturated with substrate, could maximally support an intracellular H202 production of 9.4 pmol . min * mg protein.-’ Thus, XOgenerated H202, if it exclusively diffused from cells, would account for only 13% of the basal rate of extracellular H202 release of 75 + 12 pmol * mine1 . mg protein-i in bovine aortic endothelial cells. The relative contribution of X0 to net endothelial 0; and H202 production would be much less than this proportion, when intracellular target molecule reactions of H202 and antioxidant enzyme scavenging processes are considered as well. We were unable to detect any direct effect of TNF or LPS on H202 metabolism or XDH + X0 activities. Since exposure conditions were identical to our previous studies showing cytokine-mediated loss of cell monolayer barrier function, we conclude that increased endogenous 0, and H202 production does not mediate direct TNF and LPS toxicity to bovine aortic endothelial cells. Our discrepancies with recently published reports suggesting a role for X0 in inflammatory mediator effects could involve a number of explanations, including endothelial cell heterogeneity. Because of this heterogeneity, results of studies concerning reactive oxygen species metabolism in endothelial cells from different species, from different vessels, and when grown in different culture conditions

INFLAMMATORY

MEDIATORS

AND

ENDOTHELIAL

should be interpreted in the context of that specific endothelial cell type. A future challenge is to develop an endothelial cell model which most accurately reflects physiologic and pathologic events in the human vasculature. Because of the potent effect which inflammatory cell-derived cytokines have on cell * NO production, it is possible that LPS and TNF-a may stimulate endothelial production of ONOO-, which has a reactivity similar to . OH and can participate in oxidation and nitration reactions (12, 13). ACKNOWLEDGMENTS This research was supported by American Lung Association Research Grant N-119, NM Grant NS 24275, and the Council for Tobacco Research. We are grateful to Drs. Joseph Beckman and Rafael Radi for helpful critiques and to Yvonne Lambott for manuscript preparation.

REFERENCES 1. Brigham, K. L., Meyrick, B., Berry, L. C., and Repine, J. E. (1987) J. Appl. Physiol. 63, 840-850. 2. Phan, S. M., Gannon, D. E., Varani, J., Ryan, U. S., and Ward, P. A. (1989) Am. J. Pathol. 134, 1201-1211. 3. Friedl, H. P., Till, G. O., Ryan, U. S., and Ward, P. A. (1989) FASEB J. 3,2512-2518. 4. Weiss, S. J., Young, J., LoBuglio, A. F., Slivka, A., and Nimeh, N. F. (1981) J. Clin. Invest. 68, 712-721. 5. Vercellotti, G. M., Dobson, M., Schorer, A. E., and Moldow, C. F. (1988) PFOC.Sot. Exp. Biol. Med. 187, 181-189. 6. Meyrick, B. O., Ryan, U. S., and Brigham, K. L. (1986) Am. J. Physiol. 122,140-151. 7. Beutler, B., and Cerami, A. (1988) Annu. Reu. Biochem. 57, 505518. 8. Royall, J. A., Berkow, R. L., Beckman, J. S., Cunningham, M. K., Matalon, S. M., and Freeman, B. A. (1989) Am. J. Physiol. 257 (Lung Cell. Mol. Physiol. l), L399-L410. 9. Pohlman, T. H., Stanness, K. A., Beatty, P. G., Ochs, H. D., and Harlan, J. M. (1986) J. Immunol. 136, 4548-4553. 10. Varani, J., Bendelow, M. J., Sealey, D. E., Kunkel, S. L., Gannon, D. E., Ryan, U. S., and Ward, P. A. (1988) Lab. Invest. 59, 292295. 11. Gannon, D. E., Varani, J., Phan, S. M., Ward, J. H., Kaplan, J., Till, G. O., Simon, R. H., Ryan, U. S., and Ward, P. A. (1987) Lab. Invest. 57, 37-44. 12. Beckman, J. S., Beckman, T. W., Chen, J., Marshall, P. A., and Freeman, B. A. (1990) Proc. Natl. Acad. Sci. USA 87, 1620-1624. 13. Radi, R., Beckman, J. S., and Freeman, B. A. (1991) J. Biol. Chem.

266,4244-4250. 14. Marshall,

P. J., and Landis, W. E. M. (1986) J. Lab. Clin. Med.

108,525-534. 15. Matsubara, T., and Ziff, M. (1986) J. Zmmunol. 137, 3295-3298. 16. Gorog, P., Pearson, J. D., and Kakkar, V. V. (1988) Atherosclerosis

72,19-27. 17. Jones, D. P., and Kennedy, F. G. (1983) in Functions of Glutathione: Biochemical, (Larsson, A., pp. 1099116, 18. Dobrina, A., 19. Kuppusamy, 9884.

Physiological, Toxicological, and Clinical Aspects Holmgren, A., Orrenius, S., and Manneervik, B., Eds.), Raven Press, New York. and Patriarca, P. (1986) J. Clin. Invest. 78, 462-471. P., and Zweier, J. L. (1989) J. Biol. Chem. 264,9880-

OXIDANT

METABOLISM

693

20. Bishop, C. T., Mirza, Z., Crapo, J. D., and Freeman, B. A. (1985) In Vitro Cell. Dev. Biol. 21, 229-236. 21. Cohen, C., and Hochstein, P. (1965) J. Pharmacol. Exp. Ther. 147, 139-143.

22. Bergmeyer, H. U. (1963) Methods of Enzymatic 23. 24. 25. 26. 27. 28. 29.

30. 31. 32. 33. 34. 35. 36. 37. ?Q “o. 39. 40.

Analysis, pp. 885888, Academic Press, New York. Lowry, 0. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265-275. Labarca, C., and Paigen, K. (1980) Anal. Biochem. 102, 344-352. Yusa, T., Beckman, J. S., Crapo, J. D., and Freeman, B. A. (1987) J. Appl. Physiol. 63, 353-358. Hyslop, P. A., and Sklar, L. A. (1984) Anal. B&hem. 141, 280-286. Katz, J., and Wood, H. G. (1963) J. Biol. Chem. 238, 517-523. Paglia, D. E., and Valentine, W. E. (1967) J. Lab. Clin. Med. 70, 158-169. Flohe, L., and Brand, I. (1969) B&him. Biophys. Actu 191, 541-549. Beckman, J. S., Parks, D. A., Pearson, J. D., Marshall, P. A., and Freeman, B. A. (1989) Free Radicals Biol. Med. 6, 607-615. Kono, Y., and Fridovich, I. (1982) J. Biol. Chem. 257,5751-5754. Oshino, N., Chance, B., Sies, H., and Bucher, T. (1973) Arch. Biochem. Biophys. 154, 117-131. Crapo, J. D., Freeman, B. A., Barry, B. E., Turrens, J. F., and Young, S. L. (1983) Physiologist 26, 170-176. Freeman, B. A., and Crapo, J. D. (1981) J. Biol. Chem. 256,10,98610,992. Turrens, J. F., Freeman, B. A., Levitt, J. G., and Crapo, J. D. (1982). Arch. Biochem. Biophys. 217, 401-410. Jones, D. P., Eklow, L., Thor, H., and Orrenius, S. (1981) Arch. Biochem. Biophys. 210, 505-516. May, J. M. (1981) Arch. Biochem. Biophys. 207, 117-127. Chance, B., Sies, H., and Boveris, A. (1979) Phystil. Rev. 59,527-605. O’Neil, J. J., and Tierney, D. F. (1974) Am. J Physiol. 226,867-873. Katz, J., Landau, B. R., and Bartsch, G. E. (1966) J. Biol. Chem.

241,727-740. 41. Harlan, J. M., Levine, J. D., Callahan, K. S., Schwartz, B. R., and Harker, L. A. (1984) J. Clin. Invest. 73, 706.-713. 42. Ody, C., and Junod, A. F. (1985) Proc. Sot. Exp. Biol. Med. 180, 103-111. 43. Hampel, G., Watanabe, K., Weksler, B. B., and Jaffe, E. A. (1989) Biochim. Biophys. Acta 1006, 151-158. 44. Hart, D. H. L., Hobson, J. E., Walker, D. C., and Autor, A. P. (1986) J. Free Radicals Biol. Med. 1, 429-435. 45. Shiki, Y., Meyrick, B. O., Brigham, K. L., and Burr, I. M. (1987) Am. J. Physiol. 252, C436-C440. 46. Suttorp, N., Toepper, W., and Roka, L. (1986) Am. J. Physiol. 251,

C671LC680. 47. Steinberg, P., Schramm,

H., Schladt, L., Robertson, L. W., Thomas, H., and Oesch, F. (1989) Biochem. J. 264, 737-744. 48. Lawerence, R. A., and Burk, R. F. (1976) Biochem. Biophys. Res. Commun. 71,952-958.

49. Parks, D. A., and Granger. D. N. (1986) Acta Physiol. Stand. Suppl.

648,87-99. 50. Rinaldo, J. E., and Gorry, M. (1990) Am. J. Respir. Cell Mol. Biol. 3,525-533. 51. Jarasch, E.-D., Bruder, G., and Heid, H. W. (1986) Acta Physiol. Stand. Suppl. 548, 39-46. 52. Bruder, G., Heid, H. W., Jarasch, E.-D., and Mather, I. H. (1983) Differentiation 23, 218-225. 53. Panus, P. C., Wilborn, A., Cunningham, M. K., Wright, S. W., and Freeman, B. A. (1990) Am. Rev. Respir. Dis. 114, A351.

694

ROYALL MINIPRINT

When AT concentration inactivation

is sufficiently

ET AL. SUPPLEMENT

high and not rate limiting, the rate of catalase

Because the concentration

isdescribed by:

of aminotriazole

Hz02 at the site of interaction

is high and the local concentration

with catalase in the peroxisome will be maintained

of at

a low level, then kAT [AT] > > k2 [H202] and:

d[CATl __ = -kAT [AT] [Compound I] dt

[Compound

II =

kl [CAT1 IH2021

Eq.4

kAT[ATl

Catalase concentration inactivation

by AT, [AT] is AT concentration,

concentration

of Compound

other electron determined remaining

is [CAT]. kAT is the second order rate constant for catalase

donors,

and [Compound

I. Assuming no competing

the steady state concentration

by its rate of formation

Substituting d[CATl

II is the steady state

reactions with’alcohols of Compound

and cts rate of degradation

or

I will be

by the two

Eq. 4 into Eq. 1: kl [CAT1 [Hz021 = -kArlAT] X

dt

= -k, ICAT] IH2021

Eq. 5

kArlAT

From the experimentally-derived pseudo-first

AT-mediated

catalase inactivation

order rate constant of catalase inactivation

curve, the

(kcat) can be described by:

pathways: dICAT1

d[Compound

I] = 0 = kl [CAT] [Hz021 - kAT [AT] [Compound

II

Setting Eq. 5 equal to Eq. 6:

dt - k2 [H202] [Compound II

= -k,,t [CAT]

dt

Eq. 2

-kl [CAT] [H202] = -kcat [CAT] or

The rate of Compound

I formation

with H202. By rearrangement

is k,, and k2 the rate of reaction of Compound

of Eq. 2:

kl [CAT1 W2021 [Compound

I

II =

The value of kc is known (1.7 x 107 M-1 s-1) and thus the H202 concentration Eq. 3

hiAT

+ k2W2021

calculated.

can be