Impact of pulmonary arterial endothelial cells on duroquinone redox status

Free Radical Biology & Medicine, Vol. 37, No. 1, pp. 86 – 103, 2004 Copyright D 2004 Elsevier Inc. Printed in the USA. All rights reserved 0891-5849/$-see front matter

doi:10.1016/j.freeradbiomed.2004.02.078

Original Contribution IMPACT OF PULMONARY ARTERIAL ENDOTHELIAL CELLS ON DUROQUINONE REDOX STATUS MARILYN P. MERKER,*,y,z,§ ROBERT D. BONGARD, b GARY S. KRENZ, z HONGTAO ZHAO, # VIOLA S. FERNANDES, z BALARAMAN KALYANARAMAN, # NEIL HOGG, # and SAID H. AUDI z,** *Department of Anesthesiology, y Department of Pharmacology/Toxicology, b Department of Physiology, # Department of Biophysics, **Department of Pulmonary and Critical Care Medicine, Medical College of Wisconsin, Milwaukee, WI; z Department of Biomedical Engineering and z Department of Mathematics, Statistics and Computer Science, Marquette University, Milwaukee, WI; and § Zablocki V.A. Medical Center, Milwaukee, WI 53295, USA (Received 17 December 2003; Revised 13 February 2004; Accepted 27 February 2004) Available online 26 March 2004

Abstract—The study objective was to use pulmonary arterial endothelial cells to examine kinetics and mechanisms contributing to the disposition of the quinone 2,3,5,6-tetramethyl-1,4-benzoquinone (duroquinone, DQ) observed during passage through the pulmonary circulation. The approach was to add DQ, durohydroquinone (DQH2), or DQ with the cell membrane-impermeant oxidizing agent, ferricyanide (Fe(CN)6 3), to the cell medium, and to measure the medium concentrations of substrates and products over time. Studies were carried out under control conditions and with dicumarol, to inhibit NAD(P)H:quinone oxidoreductase 1 (NQO1), or cyanide, to inhibit mitochondrial electron transport. In control cells, DQH2 appears in the extracellular medium of cells incubated with DQ, and DQ appears when the cells are incubated with DQH2. Dicumarol blocked the appearance of DQH2 when DQ was added to the cell medium, and cyanide blocked the appearance of DQ when DQH2 was added to the cell medium, suggesting that the two electron reductase NQO1 dominates DQ reduction and mitochondrial electron transport complex III is the predominant route of DQH2 oxidation. In the presence of cyanide, the addition of DQ also resulted in an increased rate of appearance of DQH2 and stimulation of cyanide-insensitive oxygen consumption. As DQH2 does not autoxidize-comproportionate over the study time course, these observations suggest a cyanide-stimulated one-electron DQ reduction and durosemiquinone S (DQ ) autoxidation. The latter processes are apparently confined to the cell interior, as the cell membrane impermeant oxidant, ferricyanide, did not inhibit the DQ-stimulated cyanide-insensitive oxygen consumption. Thus, regardless of whether DQ is reduced via a one- or two-electron reduction pathway, the net effect in the extracellular medium is the appearance of DQH2. These endothelial redox functions and their apposition to the vessel lumen are consistent with the pulmonary endothelium being an important site of DQ reduction to DQH2 observed in the lungs. D 2004 Elsevier Inc. All rights reserved. Keywords—Endothelium, Quinone, High-performance liquid chromatography, Electron paramagnetic resonance, Kinetic model, Quinone reductase, Hydroquinone oxidase, Pulmonary metabolism, Free radicals

ical composition of the plasma as the blood passes through the circulation. It does so by providing a redox active surface in apposition to the plasma capable of reducing substances from several chemical classes of redox active compounds, including quinones (e.g., menadione [1], coenzyme Q0 [2], and 2,3,5,6-tetramethyl-1,4-benzoquinone (duroquinone, subsequently referred to as DQ) [3]), thiazines (e.g., toluidine blue O polymer, methylene blue [4– 6]), viologens (e.g., paraquat [7,8]), disulfides (e.g., lipoic acid [9]), oxygen [10 – 12], mono- and dehydroas-

INTRODUCTION

Through the activities of various plasma membrane and/or intracellular transporters and enzymes, the vascular endothelium plays an important role in determining the chem-

Address correspondence to: Marilyn P. Merker, VAMC, Research Service 151, Milwaukee, WI 53295, USA; Fax: (414)-382-5374; E-mail: [email protected]. 86

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corbate [13], and probably others. Reduction affects the pro- and antioxidant activity, tissue permeability, and other properties of such compounds, making the endothelium an important determinant of their activity and disposition. The pulmonary endothelium is unique in that its surface area is nearly equal to that of the rest of the endothelium in the body, and being situated between the systemic venous and arterial circulations, it has the potential for determining the lung disposition and the bioactivity of redox active compounds passing from the venous to systemic arterial circulations. We have observed that the lungs have a large capacity for converting DQ that enters the lungs via the pulmonary arterial inflow to the reduced durohydroquinone (DQH2) form that then appears in the pulmonary venous effluent [3]. Reduction of this model amphipathic quinone appears to be predominately via NQO1, and the net DQH2 output from the lungs is dominated by the combination of NQO1-mediated reduction and DQH2 reoxidation via mitochondrial complex III. The objective of the present study is to examine whether pulmonary endothelial cells play an important role in determining the whole lung disposition of DQ and, if so, to use the cells in culture as a tool for in-depth study of the mechanisms by which the cells influence the extracellular redox status of this model quinone substrate. Of the quinone reductases identified in various cell types, including endothelial cells, NQO1 is among the most thoroughly studied [14 – 18]. It is a cytoplasmic NAD(P)H-quinone oxidoreductase that carries out two electron quinone reduction, at least one function of which is thought to be to decrease the availability of its quinoid substrates to single electron quinone reductases (e.g., cytochrome c, b5, and P450 reductases, NO synthase), thereby avoiding semiquinone generated oxidative stress [16 –20]. Since DQ reduction was shown to be predominately via NQO1 in the lung studies [3], a key goal of the present studies was to evaluate the role of NQO1 in DQ reduction by the pulmonary arterial endothelial cells. The main approach taken was to add DQ, DQH2, or DQ along with the cell membrane-impermeant redox indicator, ferricyanide (Fe(CN)6 3), to the extracellular medium under control conditions and in the presence of the NQO1 inhibitor, dicumarol, and the mitochondrial electron transport inhibitor, cyanide. The medium concentrations of the substrates and products, and cellular oxygen consumption, were measured over time. In addition, NQO1 immunoblotting was used to identify this enzyme in the cells. The hypotheses developed to explain the observations were expressed in the form of a kinetic model. The data and the model were used to evaluate the contributions of NQO1 and other pulmonary arterial endothelial cell redox pathways to the net effect of the

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cells on DQ redox status and to their role in the whole lung disposition of DQ. MATERIALS AND METHODS

Duroquinone (2,3,5,6-tetramethyl-1,4-benzoquinone, DQ), potassium hexocyanoferrate(III) (Fe(CN)6 3, ferricyanide), N-2-hydroxyethylpiperazine-N V-2-ethanesulfonic acid (Hepes), and Lactate Dehydrogenase Assay Kit 340-LD were purchased from Sigma Chemical Company (St. Louis, MO, USA). RPMI-1640 tissue culture medium was from Invitrogen (Carlsbad, CA, USA). Fetal bovine serum was from Hyclone Laboratories (Logan, UT, USA), and Biosilon beads were from Nunc (Roskilde, Denmark). Protein determinations were performed using the Bio-Rad protein assay reagent (Bio-Rad Laboratories, Hercules, CA, USA). Linoleic acid was purchased from Nu-Chek Prep (Elysian, MN, USA), and 2,2 V-azobis(2-methylpropionamidine) dihydrochloride was from Aldrich Chemical Company (Milwaukee, WI, USA). HPLC-grade methanol was obtained from Burdick and Jackson (Muskegon, MI, USA). Durohydroquinone (DQH2) was prepared by reduction of DQ with potassium borohydride as described in [21]. 5-tertButoxycarbonyl 5-methyl-1-pyrroline N-oxide (BMPO) was synthesized as described previously [22]. The NQO1 inhibitor ES936 was the kind gift of Dr. David Siegel (School of Pharmacy, University of Colorado Health Sciences Center). Other chemicals were purchased from Sigma Chemical Company. Endothelial cell culture Bovine pulmonary arterial endothelial cells (BPAE) were isolated from segments of calf pulmonary artery, characterized, and cultured on Biosilon microcarrier beads (mean diameter 230 Am, surface area 255 cm2/gm beads) in RPMI-1640 medium supplemented with 10% fetal calf serum, 100 U/ml penicillin, 100 Ag/ml streptomycin, and 30 mg/ml L-glutamine as previously described [5]. Cells were used between passages 4 and 20. Protocols for measuring DQ reduction and DQH2 oxidation by the intact cells Approximately 0.3 ml packed volume of cell-coated beads (equivalent to 48.8 F 8.6 (SD) cm2 cell surface area, n = 108) were aliquoted from the stirred culture flasks into 55  10  10 mm acrylic spectrophotometric cuvettes (Sarstedt, Newton, NC, USA). After the cellcoated beads had settled, they were washed three consecutive times by resuspension in 3 ml of Hanks balanced salt solution containing 5.5 mM glucose and 10 mM Hepes (HBSS/Hepes), allowing the beads to settle between each wash. The HBSS/Hepes was the experimental medium in all experiments that follow.

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The cell-coated beads were then resuspended in HBSS/Hepes containing DQ or DQH2 (50 AM), and allowed to settle below the level of the spectrophotometer light path. The absorption spectrum of the medium was measured between 250 and 350 nm using a Beckman Model DU 7400 spectrophotometer (Beckman Instruments). The cuvettes were placed on a Nutator mixer in a 37jC incubator, and periodically, the mixing was stopped and the cell-coated beads were allowed to settle to measure the medium absorption spectrum. The procedure of allowing the beads to settle prior to measuring the absorbance spectrum was repeated at intervals. After 30 min mixing time, a sample was obtained for HPLC analysis. The remaining medium was removed from the cells and H2O2 (0.1 mM, final concentration) and peroxidase (1.48 U/ml, final concentration) were added to this cell-free medium to oxidize any DQH2. The absorption spectrum was measured again to determine the total concentration of DQ present. The samples destined for HPLC analysis were diluted 1:10 into the HPLC mobile phase and injected onto the HPLC system (described below). In other experiments with this protocol, inhibitors including potassium cyanide (2 mM; cyanide), dicumarol (25 AM), cyanide plus dicumarol (2 mM and 25 AM, respectively), or ES936 (0.5 AM) were added to the medium along with the DQ or DQH2. The same protocol was also carried out with the cuvettes only, without cells present, to control for nonspecific association of DQ with the plasticware. For the data obtained spectrophotometrically, the concentration of DQ in the extracellular medium was calculated from the absorbance values at 265 nm using the extinction coefficient of 21.64 mM1 cm1, following correction for crossover absorbance from DQH2 (absorbance at 299 nm). Cell-mediated reduction of ferricyanide by DQ was also measured using a similar protocol. In that case, the HBSS/Hepes added to the cells contained DQ and ferricyanide (1– 50 and 600 AM, respectively), and the absorbance of the medium was measured at 421 nm. The concentration of ferrocyanide (Fe(CN)6 4) produced was calculated from the decrease in ferricyanide concentration determined using an extinction coefficient of 1.0 mM1 cm1. The ferricyanide protocol included experiments in which cyanide (2 mM) or dicumarol (25 AM), cyanide plus dicumarol (2 mM and 25 AM, respectively), or ES936 (0.5 AM) was added to the medium. Background ferricyanide reduction was measured under each experimental condition with no DQ added to the medium. DQ/DQH2 HPLC method The HPLC system and separation method for determination of DQ and DQH2 were as described for coenzyme Q0 [2], with modifications to the mobile phase. DQ and DQH2 were separated using a Supelcosil octadecylsilane

LC-18-T (3 AM particle size, 150  4.6 mm) column eluted with a gradient composed of A (50 mM sodium acetate (pH 3.5) containing 0.1% trifluoroacetic acid and 35% methanol) and B (50 mM sodium acetate (pH 3.5) containing 0.1% trifluoroacetic acid and 70% methanol), as follows: A for 0 to 10 min; change from A to B at 10 to 12 min; B for 12 to 20 min; change from B to A at 20 to 22 min; reequilibration in A at 22 to 30 min. The mobile phase was continuously sparged with N2 throughout the course of the HPLC studies. The DQ in the column eluate was measured by absorbance at 270 nm. DQH2 was detected electrochemically, with the potentials at the first and second analytical electrodes set at 250 and +500 mV, respectively. The first electrode served as a screen electrode to prevent interference from compounds that might co-elute with DQH2 but have a lower oxidation potential. The DQH2 was oxidized at the second electrode. The elution times were 17.69 F 0.04 (mean F SEM) min for DQ and 5.88 F 0.09 (mean F SEM) min for DQH2, for 13 and 7 determinations, respectively. The concentrations of DQ and DQH2 in cell medium were determined by peak area quantification against standard curves. NQO1 immunoblots NQO1 protein was studied in the cell cytosol enriched fraction obtained by lysing the cells (f0.2 ml of packed cell coated beads) by sonication (3 pulses of 15 s each with the power set to 35% of maximum using a Model 16– 850 Virsonic Cell Disrupter) on ice in 2 ml of 25 mM Tris-HCl buffer containing 250 mM sucrose and 1 AM FAD, pH 7.4. The lysate was centrifuged for 30 min at 10,000g at 4jC, and portions of the cell supernatant fractions containing 10 Ag protein or purified recombinant human NQO1 protein (0.25 ng) were subjected to electrophoresis using Invitrogen NuPAGE LDS sample buffer, a 4 – 12% gradient Nu-PAGE Bis– Tris polyacrylamide gel, and Mes – SDS running buffer, as previously described [3]. The proteins were transferred to a nitrocellulose membrane, which was incubated for 1 h in Tris-buffered saline containing 0.1% Tween 20 and 2% bovine serum albumin, the latter as a blocking agent. The membrane was then incubated sequentially in a 1:10 dilution of a-NQO1 monoclonal antibody from IgG1-secreting hybridomas (1:1 mixture of clones A180 and mAb B771; gift of Dr. David Siegel, School of Pharmacy, University of Colorado Health Sciences Center), a 1:7500 dilution of goat a-mouse IgG horseradish peroxidase (Jackson ImmunoResearch Laboratories), and the Supersignal West Pico Chemiluminescent substrate (Pierce). The signal was captured on CLXposure Film (Pierce). As a control, nonspecific mouse IgG1 was substituted for the a-NQO1 monoclonal antibodies.

Endothelium and quinone redox status

Oximetry Cellular oxygen consumption was measured using a Yellow Springs Instruments Model 5300 biological oxygen monitor (YSI Instrument Co., Inc., Yellow Springs, OH, USA). Cell coated beads along with 3 ml airsaturated HBSS/Hepes were placed in a sealed magnetically stirred chamber at 37jC. Additions of cyanide, DQ, DQH2 and ferricyanide to the sample chamber were made using a syringe (Hamilton Company, Reno, NV, USA). Pyridine nucleotide HPLC The cell-coated beads (f0.2 ml packed volume) were aliquoted from the stirred culture flasks into conical bottomed tubes, and after the beads settled, they were washed four times by resuspension in 3 ml HBSS/Hepes, allowing the beads to settle between washes. The cellcoated beads were then resuspended in HBSS/Hepes alone (control) or HBSS/Hepes containing 2 mM cyanide and incubated by mixing on a Nutator mixer at 37jC for 30 min. HPLC measurements of total KOH-extractable intracellular NAD+, NADH, NADP+, and NADPH were carried out as described previously after removal of the incubation medium from the cell-coated beads [23]. Linoleic acid oxidation assay The linoleic acid oxidation assay was carried out using a 16 mM linoleic acid stock solution prepared in 50 mM borate buffer containing 20 mM NaOH and 0.5% Tween 20 and stored under N2 at 4jC, as described by Liegeois et al. [24]. The linoleic acid stock solution was diluted to a final concentration of 160 AM in either HBSS/Hepes medium removed from cells that had been incubated in medium containing DQ (20 AM), HBSS/Hepes medium removed from cells incubated in medium without DQ to obtain control cell conditioned medium, or control cell conditioned medium to which DQ was added after the medium had been removed from the cells. The incubations were carried out under anaerobic conditions, obtained by bubbling of the HBSS/Hepes surrounding the cells with N2, to minimize loss of cell-generated DQH2 via cell-mediated oxidation. The solutions containing the various media and linoleic acid were placed in quartz spectrophotometric cuvettes. The linoleic acid oxidation reaction was initiated by the addition of 2,2 Vazobis(2-methylpropionamidine) dihydrochloride (AMPH; final concentration 1 mM), the contents of the cuvettes were mixed, an initial spectophotometric measurement was taken at 234 nm for determination of conjugated dienes, and a portion of the reaction was removed, diluted into HPLC mobile phase, and injected onto the HPLC system for determination of DQ and DQH2. The reactions were incubated at 37jC, and peri-

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odically, spectrophotometric and HPLC determinations were repeated. Electron paramagnetic resonance (EPR) spectroscopy The protocol for collecting samples for EPR analysis was similar to the protocol described above for the spectrophotometric determination of DQ reduction by the cells except that 100 AM DQ or menadione was added to the cell incubation medium (HBSS/Hepes), which also contained 100 AM DTPA and 50 mM of the cyclic nitrone spin trap BMPO [22]. The mixtures were incubated for 10 min, after which time the cell coated beads were allowed to settle and a portion of the medium was centrifuged at 5000g for 1 min and the supernatant transferred to a Bruker Aqua X EPR liquid sample cell. EPR spectra were recorded at room temperature on a Bruker EMX spectrometer operating at 9.85 GHz and a cavity equipped with the Bruker Aqua X liquid sample cell. Typical EPR spectrometer parameters were: scan range, 100 G; field set, 3500 G; time constant, 5.12 ms; scan time, 5.12 s; modulation amplitude, 1.0 G; modulation frequency, 100 kHz; receiver gain, 6.32  105; microwave power, 10.0 mW; and number of scans per sample, 50. The EPR spectra were simulated using the software developed by Dr. Duling from the laboratory of Molecular Biophysics, NIEHS (Research Triangle Park, NC, USA) [25]. Assay for DQ and coenzyme Q0 reduction in the presence of plasma membrane protein-coated avidin beads Plasma membrane protein coated avidin beads were prepared using a simplification of a previously described procedure [2]. Approximately 3 ml of cell-coated beads (culture area, 480 cm2) were removed from the culture bottles and washed three times by resuspension and settling in 8 ml ice-cold HBSS for each wash. Biotin labeling of the plasma membrane proteins was initiated by resuspension of the cell-coated beads in 10 ml of icecold HBSS containing 11.7 mg of sulfo-NHS-LC-biotin (Pierce Biotechnology, Inc., Rockford, IL, USA; final concentration 2.1 mM). After 1 min of mixing with intermittent placement in ice, 1 ml of ice-cold HBSS containing 10 mM glycine was added to the tube to scavenge unreacted sulfo-NHS-LC-biotin. The cell-coated beads were allowed to settle on ice, the supernatant was removed, and the cell-coated beads were washed free of the biotin labeling solution by resuspension and settling once in 10 ml ice-cold HBSS containing 10 mM glycine and twice in 10 ml each time with HBSS/Hepes. After the final wash solution was removed, the cells were solubilized in 3 ml of HBSS/Hepes containing 2% 3-[(3cholamidopropyl)dimethylammonio]-1-propanesulfonate, 3 mM EDTA, and 1 mM phenylmethylsulfonyl fluoride, with mixing at 4jC. After 60 min of mixing,

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any whole cells or beads were removed by centrifugation at 1800g for 10 min. The supernatant was removed and centrifuged at 145,000g for 45 min, and the final supernatant was then used in the preparation of plasma membrane protein coated avidin beads, as previously described [2]. DQ or coenzyme Q0 reduction in the presence of the plasma membrane-coated avidin beads was measured as previously described [2] by mixing DQ (100 AM) or coenzyme Q0 (100 AM) and NADH (100 AM) or NADPH (100 AM) with the avidin beads in HBSS/Hepes. Quinone reduction was measured as the initial rate of oxidation of NADH to NAD+ or NADPH to NADP+, respectively, as determined spectrophotometrically by absorbance at 340 nm. The background data obtained from incubating DQ or coenzyme Q0 and NADH or NADPH with avidin beads prepared from cells that were not exposed to the biotin labeling reagent were subtracted from the data obtained in the presence of avidin beads before calculating the rates of NAD(P)H oxidation. One experiment was carried out. Additional measurements As a measure of cell viability, lactate dehydrogenase (LDH) activity in the medium of cells exposed to DQ and DQH2, with and without inhibitors and with and without ferricyanide present in the medium, was determined, as previously described [5]. After the extracellular medium was removed from the cells, the cells were lysed by sonication, and LDH activity in the extracellular medium and the lysed cell fraction was measured. The fraction of the total cell LDH released into the medium for cells incubated with DQ or DQH2 was 3.0 F 0.3% (mean F SEM) for 108 experiments with control cells with or without ferricyanide and inhibitor treated cells with or without ferricyanide. No significant differences in LDH release between control and inhibitor treated cells were detected. To normalize the data for comparisons between studies, the protein content of the cells in each intact cell experiment and of the cell lysate for the avidin bead studies was measured using the Bio-Rad protein assay. The protein assay for the cells was carried out following sonication of the cells, as previously described [5], and the washed microcarrier beads were dried and weighed. The total protein per unit microcarrier bead surface area was 38.3 F 1.4 Ag/cm2 (mean F SEM) for all experiments combined, with no detectable significant differences between control and treated cells. RESULTS

Figure 1A shows the time course of DQ disappearance from the medium when DQ was added under control conditions. The DQ disappearance rate decreased with time such that the DQ medium concentration reached a

Fig. 1. Effect of cells on extracellular redox status of DQ or DQH2. (A) Concentration of DQ in the medium surrounding the cells after addition of 50 AM DQ or DQH2 to the medium. (B) After 30 min of incubation, the cells were removed from the medium, peroxidase and H2O2 were added, and the concentration of DQ was measured. The reappearance of DQ, indicated by the arrows, reflects the peroxidase-mediated oxidation of DQH2 that was present in the medium at the end of the 30 min incubation period with the cells in (A). The number of experiments carried out was 16 each with DQ or DQH2. (C) The ferrocyanide concentration generated by the addition of ferricyanide (600 AM) and 50 AM DQ to the cells in 22 experiments. Also shown are the data in the absence of DQ, when ferricyanide only was incubated with the cells (background). The latter was subtracted as background from the data obtained in the presence of DQ. The symbols represent the data (means F SEM).

near steady state by 30 min. Addition of DQH2 to the medium resulted in the appearance of DQ. The DQ appearance rate also decreased with time, resulting in a DQ concentration after about 10 min that was about the same as when DQ was added (Fig. 1A). The DQH2 concentration in the medium removed from cells after 30 min of incubation, measured spectrophotometrically after its oxidation by addition of H2O2 and peroxidase to the medium, was about the same regardless of whether DQ or DQH2 was initially added to the medium (Fig. 1B), and was consistent with HPLC measurements (Fig. 2B). Figure 1C shows results of studies using the cell membrane impermeant electron acceptor ferricyanide (Fe(CN)63), which acts as a secondary electron acceptor for measuring the unidirectional rate of DQH2 appearance in the medium. Ferricyanide itself was only very slowly reduced by the cells if DQ was not present (Fig. 1C).

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Fig. 3. Incubation of DQ or DQH2 in medium in the absence of cells. (A) DQ (5, 10, 25, and 50 AM) or DQH2 (50 AM) was incubated in the medium in spectrophotometric cuvettes, and after 30 min of incubation (B) peroxidase and H2O2 were added to oxidize any DQH2. The solid line is the model fit (Eq. (16)) fit to data.

Fig. 2. HPLC chromatograms showing (A) DQH2 or DQ standards (50 AM each); (B) 10.2 AM DQH2 and 21.1 AM DQ present in the cell medium following a 30 min incubation with 50 AM DQ; (C) 30 AM DQ and no detectable DQH2 in the cell medium following a 30 min incubation with DQ (50 AM) and ferricyanide (600 AM). The scale of the two detector signals is such that 2.6 times the DQ area = DQH2 area at equimolar concentrations.

When DQ was added to the extracellular medium along with ferricyanide, ferricyanide reduction to ferrocyanide (Fe(CN) 6 4) was observed, and the reduction rate appeared to follow zero order kinetics (Fig. 1C). In the presence of ferricyanide, no DQH2 was detected in the medium by HPLC, indicating direct reduction of ferricyanide by DQH2 (Fig. 2C). Recovery of DQ or DQH2 in the cell medium at the end of the 30 min incubations was less than the initial concentration measured in the cell medium (Figs. 1B and 2B). To evaluate the contribution of the cells to this loss, DQ and DQH2 were incubated in medium without cells present, but otherwise under the same mixing conditions as in Fig. 1. Figure 3A shows that DQ disappeared from the medium, and that the rate of disappearance was independent of DQ concentration. That the disappearance was attributable to nonspecific interactions of DQ with the spectrophotometric cuvettes, and not reduction to DQH2, is shown by the fact that there was no detectable DQH2 present in the medium after 30 min of incubation (Fig. 3B). When DQ was incubated in the cuvettes without mixing, there was virtually no loss of DQ from medium (data not shown), indicating that the disappearance of the DQ from the mixed cuvette incubations was not a result of DQ instability. Figure 3A also shows that over the 30 min incubation period, DQH2 did not autoxidize and all of the DQH2 initially added to the

medium was recovered, as detected after oxidation to DQ (Fig. 3B), indicating that DQH2 did not undergo nonspecific interactions with the cuvettes. The capacity of the cells for reducing DQ is indicated by the DQ mediated ferricyanide reduction rate obtained over the range of DQ concentrations in Fig. 4. The maximum DQ mediated ferricyanide reduction rate was 0.56 nmol min1 per cm2 cell surface area, and the DQ concentration at half the maximum rate was 1.2 AM. The data in Fig. 5 were obtained using the same protocols as in Fig. 1, except that the NQO1 inhibitors,

Fig. 4. Dose response for DQ mediated ferricyanide reduction by cells. Ferricyanide reduction rate was measured as the rate of appearance of ferrocyanide after the addition of 600 AM ferricyanide and DQ at concentrations ranging from 0 to 50 AM to the medium surrounding the cells. The reactions were carried out for 30 min, the ferricyanide-only (0 AM DQ) data were subtracted from the data obtained in the presence of the different DQ concentrations, and the DQ-mediated ferricyanide reduction rates were obtained by linear regression of the ferricyanide concentration-versus-time curves. The symbols represent the data (means F SEM, n = 4) and the line represents Eq. (17) fit to the data.

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Fig. 5. Effects of inhibitors on the extracellular redox status of DQ and DQH2 and on DQ-mediated ferricyanide reduction. The concentration of DQ in the medium surrounding the cells after addition of 50 AM DQ or DQH2 under (A) control conditions or in the presence of (C) dicumarol (25 AM) or ES936 (0.5 AM), (E) cyanide (2 mM), and (G) dicumarol (25 AM) plus cyanide (2 mM). The protocol was as in Figs. 1A and 1B, in that after 30 min of incubation in the presence of DQ or DQH2 with the inhibitors, the medium was removed from the cells at the time represented by the vertical dashed line, peroxidase and H2O2 were added to the removed medium, and the concentration of DQ in the medium was measured (B, D, F, and H). The reappearance of DQ in the medium is a result of peroxidase-mediated DQH2 oxidation, thereby indicating the amount of DQH2 that was present in the medium at the end of the 30 min incubation period in the presence of the cells. DQ-mediated ferricyanide reduction under (I) control conditions, or in the presence of (J) dicumarol or ES936, (K) cyanide, and (L) dicumarol plus cyanide, with the inhibitor concentrations as in (C), (E) and (G). The symbols represent the data (means F SEM) and the solid lines were obtained by fitting the kinetic model to the data. For (G), (H), and (L), the dashed lines are the model prediction of the results using the parameter values obtained from fitting the model to the data in (A) through (F), (I) through (K), and Fig. 8. The numbers of experiments (n) carried out were as follows: (C, D), n = 6 with DQ and n = 7 with DQH2 for dicumarol, n = 1 each for ES936; (E, F) n = 6 with DQ and n = 3 with DQH2; (G, H) n = 5 with DQ and n = 4 with DQH2; (J) n = 14 for dicumarol, n = 1 for ES936; (K) n = 5; (L) n = 5. For (A), (B), and (I) the data and number of experiments are the same as in Fig. 1.

dicumarol or ES936 (Figs. 5C, 5D and 5J), or the mitochondrial electron transport inhibitor, cyanide (Figs. 5E, 5F and 5K), or dicumarol and cyanide together (Figs. 5G, 5H and 5L) were included in the medium. As compared with the control (Figs. 5A and 5B), dicumarol decreased and cyanide increased the rate of DQ disappearance from the medium (Figs. 5C and 5E). When DQH2 was added, dicumarol increased the rate of appearance of DQ whereas cyanide blocked its appearance (Figs. 5C and 5E). At the end of 30 min of incubation, regardless of whether DQ or DQH2 was initially added to the medium, in the presence of dicumarol there was no detectable DQH2 in the medium (Fig. 5D), whereas with cyanide, virtually all of the DQH2 or DQ added to the incubation medium was present as DQH2 (Fig. 5F). The effects of the irreversible NQO1 inhibitor ES936 were similar to those of dicumarol (Figs. 5C and 5D). As

compared with the dicumarol alone (Fig. 5C), when dicumarol and cyanide were present together and DQ was added to the medium, the DQ disappearance rate increased, and when DQH2 was initially added with both inhibitors, the DQ appearance rate decreased (Fig. 5G). For the dicumarol plus cyanide condition, the medium DQ approached the same concentration over the 30 min incubation period for either DQ or DQH2 addition to the medium, although the quasi-steady state was reached at a DQ concentration lower than that observed for the control cells. As for the other conditions, the amount of DQH2 present in the medium at the end of the incubation period with dicumarol plus cyanide was not significantly different whether DQ or DQH2 was incubated with the cells (Fig. 5H). Figures 5J, 5K, and 5L show the results of including the inhibitors on DQ-mediated ferricyanide reduction to

Endothelium and quinone redox status

ferrocyanide. As compared with the control (Fig. 5I), dicumarol or ES936 almost completely blocked DQmediated ferricyanide reduction (Fig. 5J). As compared with control conditions, cyanide increased the DQ-mediated ferricyanide reduction rate by about 74% (Fig. 5K), and dicumarol plus cyanide decreased it by about 26% (Fig. 5L). The inhibitory effects of dicumarol or ES936 on the appearance of DQH2 in the extracellular medium with cells incubated with DQ and on DQ-mediated ferricyanide reduction (Figs. 5C and 5J) imply a role for NQO1. The presence of NQO1 protein in the cell cytosolic fraction was confirmed by immunoblot as shown in Fig. 6. The results from the dicumarol and dicumarol plus cyanide studies in Fig. 5 suggest that cyanide stimulates a dicumarol insensitive, or non-NQO1-mediated, DQ reduction pathway. To determine the possibility that a oneelectron DQ reduction pathway is involved, the effect of DQ on cyanide insensitive oxygen consumption was measured (Fig. 7). In the presence of cyanide, DQ stimulated cellular oxygen consumption, and the rate decreased with time (Fig. 7A). On the addition of ferricyanide, the oxygen consumption rate increased and the rate was maintained throughout the time course studied (Fig. 7A). Ferricyanide alone did not have a detectable effect on cyanide-insensitive oxygen consumption (Fig. 7C). DQH2 alone also did not stimulate cyanide insensitive oxygen consumption (Fig. 7B), but when ferricyanide was present along with the DQH2, oxygen consumption was stimulated (Fig. 7B). The combination of DQH2 plus ferricyanide did not result in oxygen consumption in the absence of cells (data not shown). Figure 8 shows the mean data obtained for DQstimulated cyanide-insensitive cellular oxygen consumption in the absence or presence of ferricyanide. A potential mechanism underlying the effect of cyanide to stimulate dicumarol-insensitive DQ reduc-

Fig. 6. Immunoblot of bovine pulmonary arterial endothelial cell NQO1. Human recombinant NQO1 (hr NQO1, 0.25 ng protein) was used as a standard. The amount of cytosolic protein loaded onto the gel was 10 Ag. The arrows indicate the positions of molecular weight markers on the blot. There were no detectable corresponding bands in the same region of the blots when control, nonspecific IgG1 was used in place of a-NQO1 antibody.

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Fig. 7. Effect of DQ on cyanide insensitive cellular oxygen consumption. The initial recordings in each panel show the background oxygen consumption in the absence of the cells. The cells were then added to the oximeter chambers. After cell-mediated oxygen consumption was recorded, cyanide (2 mM) was added and then either (A) DQ (50 AM), followed by ferricyanide (600 AM); (B) DQH2 (50 AM), followed by ferricyanide (600 AM); or (C) ferricyanide (600 AM), followed by DQ (50 AM). All additions to the oximeter chambers were made at the times indicated by the arrows. The oxygen consumption rate measured for the control cells in the absence of cyanide was 0.23 F 0.02 (mean F SEM, n = 10) nmol O2 min 1 per cm2 cell surface area.

tion could involve a cyanide-induced increase in the availability of intracellular reducing equivalents for contributing reduction pathways. That cyanide increased the reducing capacity of the cells is shown in Table 1, wherein the effect of cyanide on the ratios of reduced to oxidized intracellular pyridine nucleotides (NADH/NAD+ and NADPH/NADP+) in control and cyanide treated cells was measured by HPLC. The intracellular NADH/NAD+ ratio was increased in cyanide treated as compared with control cells, but cyanide had no significant effect on the NADPH/NADP+ ratio.

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Fig. 8. Mean data for cyanide-insensitive oxygen consumption in the presence of (A) DQ (50 AM) or (B) DQ (50 AM) plus ferricyanide (600 AM). The symbols represent the means F SEM for data from three experiments and the lines are the model fit to the data. * Data point is significantly different from the preceding data point, p < .05.

The extracellular medium from cells incubated with DQ was examined for antioxidant activity resulting from DQH2 generation by using the lag time for linoleic acid conjugated diene formation in the presence of a free radical initiator (Fig. 9). DQ was added to the medium surrounding the cells, and the extracellular medium containing cell generated DQH2 was removed from the cells and added to the linoleic acid oxidation assay mixture. The DQH2 slowly converted back to DQ, as measured by HPLC, and the lag preceding the rapid phase of linoleic acid oxidation continued until the DQH2 was exhausted. The lag time in reaction mixtures that did not contain DQH2 (either cell conditioned medium removed from cells that had not been incubated with DQ (conditioned medium) or conditioned medium to which DQ was added after removal of the medium from the cells) was about four times less than in the reaction mixture containing the cell-generated DQH2. To evaluate the potential pro-oxidant impact of DQ in the cell medium, DQ was incubated with the cells in the

Fig. 9. Effect of cell-generated DQH2 on the lag time for linoleic acid oxidation. Formation of conjugated dienes (absorbance at 234 nm) was measured after mixing linoleic acid (160 AM) with the free radical initiator (AMPH, 1 mM) and (q) cell medium containing DQH2 generated by adding DQ (20 AM) to the endothelial cells, (E) cell conditioned medium (no quinone added), and (x) cell medium conditioned by the cells to which DQ (20 AM) was added after the medium had been removed from the cells. There was no detectable DQH2 in the media in the latter two conditions (E,x). Right axis and solid lines: HPLC measurements of oxidation of DQH2 ( ) to DQ (o) in the assay condition containing the cell-generated DQH2 (q).

.

S

presence of the O2  spin trap BMPO, and the extracellular medium was removed for EPR detection of radical adduct formation. The radical adduct signal observed with DQ (Fig. 10B) was much smaller than that seen with the same concentration of menadione (Fig. 10A), the latter of which was used as a positive control for stimulation of endothelial cell superoxide production [1]. Both signals were completely suppressed by SOD (shown only for menadione; Fig. 10C), consistent with O2 S being the primary trapped reactive oxygen species,

Table 1. Effect of Cyanide on Pyridine Nucleotide Redox Status Treatment

n

NADH/NAD+

NADPH/NADP+

Control Cyanide

4 4

0.08 F 0.01 0.37 F 0.05*

6.5 F 0.7 6.9 F 0.7

+

+

The values are the mean NADH/NAD or NADPH/NADP ratios F SEM. * The NADH/NAD+ ratio was significantly higher for cyanide treated as compared to control cells, p < .005 (t test).

Fig. 10. EPR spectra of extracellular medium from cells incubated with BMPO and (A) menadione (100 AM), (B) DQ (100 Am), or (C) menadione (100 AM) and superoxide disumutase (100 units/ml). The spectrum in (D) is for BMPO with cells only. Also shown is the spectrum obtained when BMPO was incubated with menadione (100 AM) in the absence of cells (E).

Endothelium and quinone redox status Table 2. Plasma Membrane Protein-Coated Avidin Bead-Mediated DQ and Coenzyme Q0 Reduction +

NADP

V max3; K m3 1 DQ þ RH ! DQ 2

(pmol formed/min/mg beads) 45 844

85 124

2DQ

DQ and coenzyme Q0 reduction rates were measured as the initial rate of appearance of NAD+ or NADP+ resulting from NADH or NADPH oxidation, respectively.

with subsequent cellular reduction of the BMPO-OOH radical adduct to the observed adduct, BMPO-OH [22]. As transplasma membrane electron transport has been shown to play a role in pulmonary arterial endothelial cell-mediated reduction of another quinone, coenzyme Q0 [2], its potential role in DQ reduction was examined. Plasma membrane protein-coated avidin beads were incubated with DQ or coenzyme Q0, the latter as a positive control, with either NADH or NADPH as electron donors. NAD+ or NADP+ production resulting from DQ or coenzyme Q0 reduction was used to obtain a measure of the reaction rates. The initial reaction rates for DQmediated NADH or NADPH oxidation were within the same order of magnitude as for coenzyme Q0-mediated NADPH oxidation. However, the coenzyme Q0-mediated NADH oxidation rate was 10 times faster than that observed for DQ-mediated NADPH oxidation, which was the faster rate observed with DQ (Table 2). DATA ANALYSIS

A kinetic model was developed to evaluate the individual contributions of cellular processes to disposition of DQ incubated with cells (Figs. 3, 4, 5, and 8). The model includes two regions representing the cells and the medium, with volumes Vc and Vm, respectively. The model allows for NQO1-mediated two electron DQ reduction to DQH2 (Eq. (1)), cyanide-sensitive DQH2 oxidation to DQ (Eq. (2)), DQ reduction to durosemiS S quinone (DQ ) (Eq. (3)), autoxidation of DQ  to DQ S  with superoxide (O2 ) as by-product followed by disS mutation of O2  to hydrogen peroxide (H2O2) (Eq. (4)), peroxidase-mediated oxidation of DQH2 to DQ (Eq. (5)), S and DQ  disproportionation to DQH2 and DQ (Eq. (6)). In addition to these intracellular reactions, the model allows for processes occurring in the medium not attributable to cells, including DQH2-mediated ferricyanide (Fe(CN)6 3) reduction to ferrocyanide (Fe(CN)6 4) (Eq. (7)) and non-specific interactions of DQ, but not DQH2, with the spectrophotometric cuvettes (Eq. (8)). DQ þ RH þ Hþ

V max1; K m1

!

V max2; K m2 1 DQH2 þ O2 ! H2 O þ DQ 2

ð2Þ

+

NAD

DQ Coenzyme Q0

95

DQH2 þ Rþ

ð1Þ

S



S



1 1 þ Rþ þ Hþ 2 2

k1

þ O2 þ 2Hþ ! H2 O2 þ 2DQ k2

DQH2 þ H2 O2 ! DQ þ 2H2 O 2DQ

S



ð3Þ ð4Þ ð5Þ

k3

þ 2Hþ ! DQH2 þ DQ

ð6Þ

k5

2FeðCNÞ6 3 þ DQH2 ! 2FeðCNÞ6 4 þ DQ þ 2Hþ ð7Þ k6

ð8Þ

DQ þ C V DQ-C; k6

where RH and R+ are the reduced and oxidized forms, respectively, of intracellular electron donor(s); and C are cuvette sites for DQ interactions. S The reduction of DQ to DQH2 and DQ  and the cyanide-sensitive oxidation of DQH2 to DQ are assumed to follow Michaelis– Menten kinetics, where for each reaction Vmax and Km represent the apparent maximum rate and Michaelis constant, respectively. All other reactions are assumed to follow the law of mass action and to proceed with a rate constant ki in the forward direction and, if reversible within the time course of the study, with a rate constant ki in the reverse direction. DQ and DQH2 are assumed to have free access to the regions representing the cells and the medium [3]; i.e., the concentrations of DQ and DQH2 at the intracellular reaction sites are essentially in equilibrium with their respective medium concentrations. The same is assumed S for H2O2 and O2. DQ  is assumed to be confined to the region representing the cells as revealed by the results in Figs. 7 and 8. Using mass balance and mass action, the temporal variations in the concentrations of the various species in medium or cell region are described by Eqs. (9) – (15).  d½DQ

1 V max1 ½DQ

V max2 ½DQH2

¼ þ  dt Vm K m1 þ ½DQ K m2 þ ½DQH2

 V max3 ½DQ

2 ¯  k1 ½O2 ðDQ  Þ2 þ K m3 þ ½DQ

Vm k¯ 3 þ k2 ½DQH2 ½H2 O2 þ ðDQ  Þ2 Vm þ k6 ½DQ-C  k¯ 6 ½DQ

S

S

þ k5 ½DQH2 ½FeðCNÞ6 3 2

ð9Þ

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  d½DQH2

1 V max1 ½DQ

V max2 ½DQH2

¼  dt V m K m1 þ ½DQ K m2 þ ½DQH2

k¯ 3  k2 ½DQH2 ½H2 O2 þ ðDQ  Þ2 Vm k5 ½DQH2 ½FeðCNÞ6 3 2 ð10Þ

S

dDQ dt

S



¼

V max3 ½DQ

 2ðDQ K m3 þ ½DQ

S Þ ðk¯ ½O þ k¯ Þ  2

1

2

3

product of O2 partial pressure and medium O2 solubility. In the presence of Fe(CN)6 3, it is assumed that oxidation of DQH2 by Fe(CN)6 3 in the medium was very rapid in comparison to the other DQH2 oxidation pathways. Estimation of model parameters The first step in fitting the model to the data in Figs. 3, 4, 5, and 8 was to obtain the parameters descriptive of the interactions of DQ with the cuvette in the absence of cells (Fig. 3). Without cells, the medium [DQ](t) is given by

ð11Þ d½H2 O2

1 ¯ ¼ k1 ½O2 ðDQ dt Vm

SÞ

 2

 k2 ½DQH2 ½H2 O2

ð12Þ

d½DQ-C ¯ ¼ k6 ½DQ  k6 ½DQ-C

dt    d½O2 1 1 V max2 ½DQH2

¼ þ k¯ 1 ½O2 ðDQ dt V m 2 K m2 þ ½DQH2

ð13Þ

S Þ  2

ð14Þ d½FeðCNÞ6 3

¼ 2k5 ½DQH2 ½FeðCNÞ6 3 2 ; dt

ð15Þ

where [DQ](t), [DQH2](t), [Fe(CN)6 3](t), and [H2O2](t) are the respective medium concentrations of DQ, DQH2, S Fe(CN)6 3, and H2O2 at time t; DQ  (t) is the amount S  of DQ in the cell region at time t; [O2](t) is the oxygen concentration in cell and medium regions; [DQ-C](t) is the concentration of DQ bound to cuvette at time t. Note that in deriving Eqs. (9) – (15), it is assumed that VmHVc [2], and that [H+] and [RH] are constant during a given sample collection period. The model parameters are: the respective apparent maximum rates of NQO1-mediated DQ reduction to DQH2, cyanide-sensitive mitochondrial DQH2 oxidation S to DQ, and DQ reduction to DQ ; their corresponding apparent Michaelis constants; the net rate constant of S S DQ  autoxidation followed by O2  dismutation to + 2 1 1 H2O2, k¯1 = k1[H ] /Vc (nmol min AM1); the rate constant of peroxidase-mediated DQH2 oxidation, k2 S (min1 AM1); the rate constant of DQ  disproportion+ 2 1 1 ation, k¯3 = k3[H ] /Vc (nmol min ); and the respective rate constants for DQ binding and unbinding to cuvette, k¯6 = k6 [C] and k6 (min1). Except for experiments involving measurement of O2 consumption (Fig. 8), [O2](t) was assumed to be constant during a given sample collection period and is fixed at 230 AM, which is the

½DQ ðtÞ 1 ¯ ¼¯ ðk6 þ k¯ 6 eðk6 þk6 Þt Þ; ½DQ 0 k6 þ k6

ð16Þ

where [DQ]0 is the initial (t = 0) [DQ] added to cell-free medium. By fitting Eq. (16) to the data in Fig. 3, each normalized to its corresponding [DQ]0, k¯6 and k6 were estimated at 0.028 and 0.051 min1, respectively. In the absence of dicumarol and cyanide, the Fe(CN)6 3 reduction rate in the presence of DQ follows zero-order kinetics with a rate equal to twice the rate of dicumarol-sensitive DQ reduction to DQH2 (see Discussion and Appendix). Thus, the Vmax1 and Km1 for the NQO1-mediated reduction pathway were obtained by fitting Eq. (17) to the DQ-mediated Fe(CN)6 3 reduction rates in Fig. 4 (see Appendix): FeðCNÞ6 3 reduction rate ¼

2V max1 ½DQ 0 : K m1 þ ½DQ 0

ð17Þ

The estimated values of Vmax1 and Km1 are given in Table 3. Assuming the NQO1-mediated two electron reduction pathway is dominant in the absence of cyanide and dicumarol, the kinetic model condenses to Eqs. (9), (10), and (13), with Vmax3, k¯1, k2, k¯3, and k5 set to zero. The estimates of Vmax2 and Km2 given in Table 3 were obtained by fitting the solution of the model to the mean data in Figs. 5A and 5B. To estimate parameters for the one-electron DQ reS duction to DQ  pathway, the assumption that the NQO1-mediated two-electron DQ reduction to DQH2 pathway is dominant was relaxed in the presence of cyanide. Figure 7 shows that cyanide significantly decreased, but did not completely inhibit, oxygen consumption. Thus, in the presence of cyanide Vmax2 was set at 4.3 nmol/min, which is the value of Vmax2 estimated from the data in Figs. 5A and 5B scaled by the ratio of O2 consumption before (12.1 F 3.0 (SD, n = 3) nmol/min) and after (1.5 F 0.4 nmol/min) cell treatment with cyanide (see Discussion). Oxygen concentration in medium and cell regions, [O2](t), was set at 230 AM and

Endothelium and quinone redox status Table 3. Parameter Values for Endothelial Cell Redox Processes Estimated from the Data in Figs. 3, 4, 5, and 8 Reaction

Symbol

Km (AM)

Symbol

Vmax (nmoles min 1 per cm2)

NQO1-mediated DQ reduction to DQH2 Mitochondrial DQH2 oxidation One-electron DQ reduction to DQ 

Km1

1.2

Vmax1

0.28

Km2

10.7

Vmax2

0.67

Km3

13.9

Vmax3

0.89

S

assumed constant for the data in Fig. 5, but allowed to change for the data in Fig. 8 wherein O2 consumption was measured. With the values of, k¯6, k6, Vmax1, Km1, and Km2 set at the values estimated from the data in Figs. 3, 4, and 5A, Vmax3, Km3, k¯1, k2, k¯3 were estimated by simultaneously fitting the solutions of Eqs. (9) –(15) to the mean data in Figs. 5 (E, F and K) and Fig. 8 with the appropriate initial conditions. The estimated values are Vmax3 = 0.89 nmol min1 per cm2 of cell surface area, Km3 = 13.9 AM, k¯1 = 0.06 nmol1 min1 AM1, k2 = 0.16 min1 AM1, and k¯3 = 20.2 nmol1 min1. The solid lines superimposed on the data in Figs. 3, 4, 5, and 8 are the model fit to the data. The dashed lines in Figs. 5G, 5H and 5L are model predictions (see Discussion). The estimated values for the key cell-mediated rate constants from the model fits to the data are given in Table 3. DISCUSSION

The present studies were carried out to evaluate the redox processes contributing to the net effect of endothelial cells on the disposition and extracellular redox status of DQ. These results demonstrate the capacity of the normal pulmonary arterial endothelial cells to convert DQ to DQH2, with the DQH2 appearing in the extracellular medium. A variety of experimental protocols, including kinetic studies of DQ reduction, DQH 2 oxidation, and DQ-mediated ferricyanide reduction with and without inhibitors (i.e., cyanide and dicumarol), together with oxygen consumption studies, were used to evaluate the contributing redox processes. A kinetic model that incorporated hypotheses regarding the roles of individual redox processes was used to interpret the data. The results are expressed in the form of a schematic diagram summarizing the roles of these processes under different experimental conditions (Fig. 11). Under control conditions (Fig. 11A), the dominant mechanism contributing to DQ reduction appears to be two-electron reduction catalyzed by NQO1, as demonstrated by the inhibitory effect of dicumarol and the irreversible NQO1 inhibitor, ES936 (Figs. 5C and 5J). That ES936 reproduced the dicumarol effect is evidence that the key effect of dicumarol under these conditions

97

was to inhibit NQO1. A role for NQO1 is also supported by the observation that the BPAE contain NQO1 protein (Fig. 6), consistent with immunohistochemical studies showing prominent staining of endothelial NQO1 in human lung [15]. The data also show that incubation of the cells with DQH2 under control conditions results in the appearance of DQ in the medium (Fig. 5A). As DQH2 does not autoxidize over the time course of the studies (Fig. 3), the result is indicative of cell-mediated DQH2 oxidation. Insight into the oxidation mechanism was provided by the observation that cyanide blocked the appearance of DQ in the medium of cells incubated with DQH2 (Fig. 5E). The effect of cyanide implied that DQH2 oxidation is mediated primarily by mitochondrial electron transport complex III, wherein cyanide inhibition of complex IV promotes complex III reduction, closing it for DQH2 oxidation [3] (Fig. 11C). This would be consistent with the concept that NQO1-mediated reduction of low-molecular-weight quinones followed by oxidation of the hydroquinones via complex III is a means of overcoming the effects of complex I deficiencies on mitochondrial ATP production [26]. The fact that DQH2 does not stimulate cyanide insensitive oxygen consumption (Fig. 7B) further suggested that the failure of DQ to appear in the medium of cyanide-treated cells incubated with DQH2 (Fig. 5E) is due predominately to inhibition of DQH2 oxidation. Thus, the dominant route of DQH2 oxidation mechanism in control cells appears to be via mitochondrial electron transport complex III (Fig. 11A). The studies with ferricyanide provided a means for separating the effects of DQ reduction from DQH2 oxidation, thereby providing additional insight into the contribution of cell-mediated DQH2 oxidation to the net effect of the cells on DQ. In control cells, the rate of DQmediated ferricyanide reduction, which is a result of DQ reduction to DQH2 and its appearance in the medium, is twice the rate of cellular production of DQH2 when DQ is added to the cells in the absence of ferricyanide (Figs. 5A and 5I). We have hypothesized that this is a consequence of the free permeation of DQH2 into and out of the cells [3] and the ability of ferricyanide to act as an extracellular oxidant sink for DQH2 formed within the cells. According to this explanation, ferricyanide competes with cell-mediated DQH2 oxidation, essentially eliminating its contribution to the net effect of the cells on the DQ (Fig. 11B). Thus, the kinetic model used to interpret the control cell data in the absence of ferricyanide represents the hypotheses that NQO1-mediated DQ reduction and complex III-mediated DQH2 oxidation are the key cellular redox processes contributing to the disposition and extracellular redox status of DQ (Fig. 11A). The ratio of the DQH2 oxidation rate to the competing NQO1mediated DQ reduction rate determines the near-steady-

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Fig. 11. Schematic diagram of the dominant redox processes involved in the appearance of extracellular DQH2 in the medium of endothelial cells incubated with DQ, under the control (A), control + ferricyanide (B), cyanide (C), and cyanide + ferricyanide (D) conditions, as identified by experimental observations and kinetic model analysis. Processes are as follows: (1) both DQ and DQH2 freely permeate the cell; (2) NQO1 catalyzes DQ reduction to DQH2, inhibited by dicumarol and ES936; (3) mitochondrial electron transport chain (complex III) catalyzes DQH2 oxidation to DQ, inhibited by cyanide; (4) unidentified one electron quinone reductase   to catalyzes DQ reduction to DQ , activated by cyanide-induced increase in intracellular NADH; (5) disproportionation of DQ  reaction with O2, which accounts for DQ mediated, cyanide insensitive oxygen consumption; (7) superoxide DQ + DQH2; (6) DQ  to H2O2 and O2; (8) peroxidase-mediated oxidation of DQH2 to DQ; (9) ferricyanide acts as a dismutase catalyzes dismutation of O2 sink for DQH2 that competes with cell-mediated oxidation processes, e.g., reactions (3) and (8). The stoichiometric forms of the reactions appear in the text. The arrow sizes are schematic, and not representative of relative reaction rates.

S

S

S

S

state concentrations of DQH2 and DQ in the cell medium attained over the 30 minute incubation period when either DQH2 or DQ is added to the cell medium (Figs. 5A and 5B). In the presence of ferricyanide, NQO1mediated DQ reduction is the dominating reaction, with no cell-mediated DQH2 oxidation (Fig. 11B). The fit of the model to the data in Figs. 5A, 5B and 5I suggests that the hypotheses are consistent with the data. The rate constants for the NQO1-mediated DQ reduction to DQH2 and complex III-mediated DQH2 oxidation to DQ indicate that the Vmax and Km for DQH2 oxidation to DQ are larger than those for DQ reduction to DQH2, the latter of which approaches zero-order kinetics at relatively low DQ concentrations (Km1 = 1.2 AM) (Table 3). The NQO1-mediated DQ reduction represents a high-capacity reducing system, which can be appreciated by the fact that the reduction rate (0.28 nmol min1 per cm2 cell surface area) is on the same order as cellular oxygen

consumption (0.23 nmol min1 per cm2 cell surface area), wherein the O2 consumption rate is multiplied by 2 for comparison with DQ reduction rate. When the pathways that dominate the net effect of the cells on DQ are inhibited, additional DQ redox pathways are revealed. For example, under the assumption that ferricyanide competes with cell-mediated DQH2 oxidation by acting as a DQH2 sink, the prediction would be that cyanide, an inhibitor of cell-mediated DQH2 oxidation, would not affect the DQ-mediated ferricyanide reduction rate. In fact, cyanide significantly increased the measured DQ-mediated ferricyanide reduction rate as compared with control, by about 60% (from 0.56 nmol min1 per cm2 cell surface area in Fig. 5I to 0.90 nmol min1 per cm2 in Fig. 5K). In other words, without an additional pathway, if Vmax1 were set in accordance with the measured DQ-mediated ferricyanide reduction rate in the presence of cyanide, the

Endothelium and quinone redox status

kinetic model would predict a cyanide-induced DQ disappearance rate almost 2-fold greater than the observed rate (Fig. 5E). Thus, kinetic model analysis suggested a contribution of additional redox processes in cyanide-treated cells, and a clue to their nature was provided by the oxygen consumption data (Figs. 7 and 8). The observation that DQ stimulated oxygen consumption in cyanide-treated cells implied a role for a one-electron DQ reduction pathway, perhaps activated by the cyanide-induced increase in the endothelial cell reducing capacity, as reflected in the NADH/NAD+ ratio S (Table 1). The reaction of the resulting DQ  with oxygen, including redox cycling, would account for the oxygen consumption (Figs. 7, 8, and 11C). DisproS portionation of a fraction of the DQ  would contribute to DQH2 generation, in addition to that provided by NQO1-mediated DQ reduction, which together would provide for an increase in the ferricyanide reduction rate as compared with control. In addition, whereas ferricyanide acts as a sink for any DQH2 formed (Fig. 11D) in its absence, a competing cyanide-insensitive intracellular DQH2 oxidation reaction is proposed as the mechanism preventing a fraction of the DQH2 from appearing in the extracellular medium. This sequence provides an explanation for the observation that the amount of DQH2 available to be released into the medium is less in the absence than in the presence of ferricyanide in the cyanide-treated cells. To accommodate the one electron reduction process, with subsequent autoxidation and disproportionation reactions, as illustrated in Figs. 11C and 11D, additional stoichiometric equations (Eqs. (3) – (6)) were incorporated into the reaction sequences. The kinetic model analysis indicates that the model hypotheses are capable of reproducing the dominant trends in the data obtained when cyanide was present (Figs. 5E, 5F and 5K). The Vmax for the one-electron DQ reduction reaction is 3-fold greater, and the Km more than an order of magnitude greater, than for the two-electron NQO1-mediated reduction reaction (Table 3). Given the preceding explanation for the data, the prediction would be that in cyanide-treated cells, oxygen consumption would be increased by DQ until the DQ was converted to DQH2, that DQH2 alone would have little effect on oxygen consumption, and that ferricyanide plus DQ or DQH2 would also increase oxygen consumption. The latter produces the apparently paradoxical situation in which adding an oxidizing agent, ferricyanide, increases rather than decreases oxygen consumption. This is because ferricyanide rapidly converts any DQH2 to DQ, thereby supplying a constant source of DQ for the one-electron reduction pathway and subsequent autoxidation. The oxygen consumption data in Figs. 7 and 8 bore out these predictions. Furthermore, if either

99

S

the one-electron DQ reduction and/or DQ  autoxidation occurred outside the cells, it might be expected that ferricyanide, which is membrane impermeant, would have blocked DQ-stimulated cyanide-insensitive oxygen S consumption by competing with oxygen for DQ  autoxidation. The fact that it did not (Fig. 7A) is evidence that one-electron DQ reduction and redox cycling are confined to the intracellular compartment. Importantly for this interpretation, the combination of DQH2 and ferricyanide, in the absence of cells, does not stimulate oxygen consumption. The kinetic model containing stoichiometric expressions for NQO1-mediated reduction to DQH2, one-elecS tron DQ reduction to DQ , and cyanide-sensitive and insensitive pathways of DQH2 oxidation was evaluated by using it to predict, or simulate, the expected outcome for the condition in which dicumarol and cyanide were present together (Figs. 5G, 5H and 5L). This was accomplished using the parameter values obtained by fitting the model to the control, dicumarol, and cyanide (Figs. 5A – 5F and 5I – 5K) data. For the simulation, because dicumarol alone almost completely inhibits the appearance of extracellular DQH2 from DQ, and cyanide increases the reductive capacity of the cells, the contribution of NQO1 was set to zero, and all the DQH2 produced in the extracellular medium was assumed to S result from one-electron reduction followed by DQ  disproportionation, and to be limited by a cyanideinsensitive DQH2 oxidation reaction. The outcome predicted by these hypotheses reproduced the dominant trends in the data, providing a validation of the model (Figs. 5G, 5H and 5L). Some steps in the reaction sequences in Fig. 11 and the stoichiometric equations are not unique explanations for the kinetic observations. Alternative and/or additional pathways that serve the same general roles may be envisioned. For example, a question raised by this study is the mechanism of DQH2 oxidation observed in the presence of dicumarol plus cyanide. The implication is that although cyanide blocks the predominating, complex III-mediated DQH2 oxidation, another, cyanide-insensitive DQH2 oxidation pathway(s) must be available. Its contribution in the presence of cyanide and absence of dicumarol is not detectable because any DQ formed by this secondary pathway is rapidly reduced via NQO1 (Fig. 5E). However, when dicumarol is also present with the cyanide, DQ reduction is suppressed, and DQ accumulation in the medium can be observed, although it occurs at a relatively slow rate (Fig. 5G). A peroxidase reaction is proposed because it is known that hydroquinones are good peroxidase substrates [27 – 29], a fact that was also taken advantage of as a tool in the present study (Figs. 1B, 3B, 5B, 5D, 5F and 5H). However, the oxidation process might also be a result of the reaction

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between DQH2 and reactive oxygen species generated when the mitochondrial electron transport chain is blocked with cyanide [30]. The key point is that to explain all the kinetic and oxygen consumption data, it appears that the following reactions are needed: NQO1mediated DQ reduction, complex III-mediated DQH2 oxidation, a cyanide-insensitive DQH2 oxidation, and a one-electron DQ reduction followed by a combination of S DQ  dismutation and autoxidation. When the control cells were incubated with DQ, there S was no detectable EPR signal for DQ  in the extracellular medium when studies were carried out under the same conditions in which a distinct coenzyme Q0 semiquinone signal was detected in a previous study [2]. This is not too surprising, as the predominant reaction contributing to DQ reduction in control cells (i.e., NQO1) produces DQH2 in the extracellular medium and DQH2 does not autoxidize-comproportionate during the time course of the studies. However, even had it been present, S DQ  might not be stable enough to detect [31]. Therefore spin trapping studies were carried out to detect any S extracellular O2  that might have been produced. The results showed that the BMPO spin adduct signal in the extracellular medium of cells incubated with DQ was much smaller than that observed for menadione. Intracellular semiquinone autoxidation in the presence of the BMPO spin trap would not necessarily be expected to give much of an extracellular EPR signal under the assumption that superoxide dismutase and catalase would compete with the spin trap for activated oxygen species. The explanation put forth in prior studies for the SODS inhibitable O2  spin adduct formation by endothelial cells exposed to menadione was that intracellular reduction was followed by diffusion of the resulting menadiol back into the medium where it encountered menadione, S resulting in comproportionation-autoxidation, and O2  production [1]. The results with DQ reveal cellular redox processes that could potentially contribute to the net outcome for any redox active compound. But for each compound, depending on its properties with respect to its propensity to undergo any of the reactions revealed by this study, the relative weight of any one class of reactions would be different. For example DQH2 autoxidation-comproportionation reactions were set to zero in this analysis, but not for previous studies with coenzyme Q0 hydroquinone, consistent with experimental observations [2]. TPMET was not considered to play a dominant role in DQ reduction based on the observations that DQ freely permeates the cells [3] and that it was a less effective substrate for the endothelial plasma membrane proteins than was coenzyme Q0 (Table 2) [2]. In addition, dicumarol, which does not inhibit at least certain plasma membrane-associated quinone reductases [32], nearly

completely inhibited DQ mediated ferricyanide reduction by the cells, whereas it resulted in only partial inhibition of coenzyme Q0 mediated ferricyanide reduction [2]. DQ and DQH2 have been used extensively in studies of quinone reductases and mitochondrial electron transport, which, consistent with the findings of the present study, have included the use of DQ as an NQO1 substrate and DQH2 as a complex III substrate [3,33 – 37]. DQ has also been considered a redox cycling, oxygen activating quinone due to its propensity to undergo one-electron reduction, produce reactive oxygen species and, consequently, exert prooxidant effects in muscle and cardiac tissue, hepatocytes, and other cell types [38 – 45]. The endothelial cells also appear to mediate one-electron DQ reduction, but the reductase was not identified other than that it is apparently sensitive to the intracellular NADH/NAD+ ratio. Candidate reductases using NADH as an electron acceptor include NAD(P)H cytochrome c reductase [44,46] and NADH-cytochrome b 5 reductase [46]. NADPH-cytochrome P450 also catalyzes one-electron DQ reduction [37], but its role under the conditions of the present study may be less important because it utilizes NADPH preferentially as the electron donor. The two-electron reduction reactions carried out by NQO1 are often considered protective, or antioxidant, due to competition with one-electron reductases that produce redox cycling semiquinones [16 – 18]. Consistent with this concept, the results of the present study suggest that NQO1-mediated DQ reduction limits the DQ available to one-electron reduction pathway. The NQO1generated DQH2 conferred protection in the linoleic acid oxidation assay, and relatively low levels of radical species were detected in the extracellular medium of cells incubated with DQ. However, for hydroquinones that are readily autooxidizable, wherein comproportionation of the quinone and hydroquinone produce redox cycling semiquinone, NQO1 activity may promote production of reactive oxygen species in the medium [16,19,47]. The potential for paradoxical NQO1 effects is exemplified by the observation that dicumarol enS hanced O2  production elicited by menadione, but inhibited O2S production elicited by the pseudomonad quinoids phthiocol and pyocyanine in human lung A549 cells [48]. These results are consistent with the pulmonary endothelium being the dominant site of DQ reduction observed in mouse and rat lungs [3]. Like the cells, the dominant processes contributing to DQ redox status in the venous effluent on passage of DQ through the lung vasculature were NQO1-mediated DQ reduction and mitochondrial electron transport complex III-mediated DQH2 oxidation. The estimated net NQO1-mediated DQ reduction rate to DQH2 in the present study under control conditions was 0.28 nmol min1 per cm2 endo-

Endothelium and quinone redox status

thelial cell surface area, which is close to those obtained for rat (0.8 nmol min1 per cm2 endothelial surface area) and mouse (0.11 nmol min1 per cm2 endothelial surface area) lungs [3]. Likewise, the estimated Km for the cell NQO1-mediated DQ reduction to DQH2 under control conditions (1.2 AM) (Table 3) is close to that obtained for rat (1.3 AM) and mouse (3.4 AM) lungs [3]. Despite the vast literature on quinone metabolism by isolated enzymes, proteins, and subcellular fractions, including mitochondria, microsomes, and cytosol [33, 34,46,49 –53], there are relatively few kinetic studies in intact cells or tissues [2,3,54 –56]. The more common experiment involves measurement of the consequences as a signature of the resulting oxygen activation [1,47,57 – 61] or semiquinone production [62,63]. Envisioning duroquinone as a model redox active compound, the kinetic approach has the capability of revealing redox pathways and allowing for estimation of parameter values for the kinetic processes involved, thereby providing for quantitative evaluation of changes in the balance between available pathways under different experimental conditions. This approach provides a tool for identification of potential therapeutic targets for manipulation of the metabolism of redox active compounds. Vascular endothelial cells, along with hepatocytes, are unique insofar as they are in direct contact with the plasma and, thus, have the opportunity to encounter bloodborne xenobiotic (e.g., redox cycling metabolites of cigarette smoke), nutritive (e.g., antioxidant polyphenolic compounds), physiological (e.g., redox cycling dopamine and other catecholamine metabolites), and pharmacological (e.g., arylating and redox cycling chemotherapeutic agents) quinones [64]. The pulmonary endothelium has the advantage over hepatocytes with respect to potential influence on the redox status, tissue distribution, and bioactivity of these compounds in the blood in that it is so large, and virtually all of the venous blood passes over it. For spent, or oxidized, forms of plasma antioxidants or co-antioxidants that permeate the pulmonary endothelium in the oxidized and reduced forms and have a propensity to undergo a two-electron NQO1-mediated reduction, wherein the reduced form is relatively stable, the pulmonary endothelium may provide a means for regeneration of their antioxidant forms in preparation for entry into the systemic circulation.

[3]

[4]

[5]

[6]

[7] [8]

[9] [10] [11] [12] [13] [14] [15] [16] [17]

[18]

[19] [20]

Acknowledgments — This work was supported by NIH Grants HL65537, HL-24349, and P41 EB001980 and the Department of Veterans Affairs. [21] REFERENCES [1] Rosen, G. M.; Freeman, B. A. Detection of superoxide generated by endothelial cells. Proc. Natl. Acad. Sci. USA 81:7269 – 7273; 1988. [2] Audi, S. H.; Zhao, H.; Bongard, R. D.; Hogg, N.; Kettenhofen, N. J.; Kalyanaraman, B.; Dawson, C. A.; Merker, M. P. Pulmonary

[22]

101

arterial endothelial cells affect the redox status of coenzyme Q0. Free Radic. Biol. Med. 34:892 – 907; 2003. Audi, S. H.; Bongard, R. D.; Dawson, C. A.; Siegel, D.; Roerig, D. L.; Merker, M. P. Duroquinone reduction during passage through the pulmonary circulation. Am. J. Physiol. Lung Cell. Mol. Physiol. 285:L1116 – L1131; 2003. Bongard, R. D.; Merker, M. P.; Shundo, R.; Okamoto, Y.; Roerig, D. L.; Linehan, J. H.; Dawson, C. A. Reduction of thiazine dyes by bovine pulmonary arterial endothelial cells in culture. Am. J. Physiol. Lung Cell. Mol. Physiol. 269:L78 – L84; 1995. Merker, M. P.; Bongard, R. D.; Linehan, J. H.; Okamoto, Y.; Vyprachticky, D.; Brantmeier, B. M.; Roerig, D. L.; Dawson, C. A. Pulmonary endothelial thiazine uptake: separation of cell surface reduction from intracellular reoxidation. Am. J. Physiol. Lung Cell. Mol. Physiol. 272:L673 – L680; 1997. Audi, S. H.; Bongard, R. D.; Okamoto, Y.; Merker, M. P.; Roerig, D. L.; Dawson, C. A. Pulmonary reduction of an intravascular redox polymer. Am. J. Physiol. Lung Cell. Mol. Physiol. 280:L1290 – L1299; 2001. Tampo, Y.; Tsukamoto, M.; Yonaha, M. Superoxide production from paraquat evoked by exogenous NADPH in pulmonary endothelial cells. Free Radic. Biol. Med. 27:588 – 595; 1999. Britigan, B. E.; Roeder, T. L.; Shasby, D. M. Insight into the nature and site of oxygen centered free radical generation by endothelial cell monolayers using a novel spin trapping technique. Blood 79:699 – 707; 1992. Jones, W.; Li, X.; Qu, Z. C.; Perriott, L.; Whitesell, R. R.; May, J. M. Uptake, recycling, and antioxidant actions of alpha-lipoic acid in endothelial cells. Free Radic. Biol. Med. 33:83 – 93; 2002. Bongard, R. D.; Roerig, D. L.; Johnston, M. R.; Dawson, C. A. Perfusate cytochrome c reduction in isolated rabbit lungs. J. Appl. Physiol. 71:1705 – 1713; 1991. Fisher, A. B.; Al Mehdi, A. B.; Wei, Z.; Song, C.; Manevich, Y. Lung ischemia: endothelial cell signaling by reactive oxygen species. A progress report. Adv. Exp. Med. Biol. 510:343 – 347; 2003. Griendling, K. K.; Sorescu, D.; Ushio-Fukai, M. NAD(P)H oxidase: role in cardiovascular disease and pathology. Circ. Res. 86:494 – 501; 2000. May, J. M.; Qu, Z.; Neel, D. R.; Li, X. Recycling of vitamin C from its oxidized forms by human endothelial cells. Biochim. Biophys. Acta 1640:153 – 161; 2003. Lind, C.; Cadenas, E.; Hochstein, P.; Ernster, L. DT-diaphorase: purification, properties, and function. Methods Enzymol. 186: 287 – 301; 1990. Siegel, D.; Ross, D. Immunodetection of NAD(P)H:quinone oxidoreductase 1 (NQO1) in human tissues. Free Radic. Biol. Med. 29:246 – 253; 2000. Cadenas, E. Antioxidant and prooxidant functions of DT-diaphorase in quinone metabolism. [review]. Biochem. Pharmacol. 49: 127 – 140; 1995. Lind, C.; Hochstein, P.; Ernster, L. DT-diaphorase as a quinone reductase: a cellular control device against semiquinone and superoxide radical formation. Arch. Biochem. Biophys. 216: 178 – 185; 1982. Joseph, P.; Long, D. J.; Klein-Szanto, A. J.; Jaiswal, A. K. Role of NAD(P)H:quinone oxidoreductase 1 (DT diaphorase) in protection against quinone toxicity. Biochem. Pharmacol. 60:207 – 214; 2000. Cadenas, E.; Hochstein, P.; Ernster, L. Pro- and antioxidant functions of quinones and quinone reductases in mammalian cells. Adv. Enzymol. Relat. Areas Mol. Biol. 65:97 – 146; 1992. Dinkova-Kostova, A. T.; Talalay, P. Persuasive evidence that quinone reductase type I (DT diaphorase) protects cells against the toxicity of electrophiles and reactive forms of oxygen. Free Radic. Biol. Med. 29:231 – 240; 2001. Cadenas, E.; Boveris, A.; Ragan, C. I.; Stoppani, A. O. M. Production of superoxide radicals and hydrogen peroxide by NADH – ubiquinone reductase and ubiquinol – cytochrome c reductase from beef-heart mitochondria. Arch. Biochem. Biophys. 180:248 – 257; 1997. Zhao, H.; Joseph, J.; Zhang, H.; Karoui, H.; Kalyanaraman, B. Synthesis and biochemical applications of a solid cyclic nitrone

102

[23]

[24]

[25] [26]

[27]

[28]

[29]

[30]

[31]

[32]

[33]

[34]

[35]

[36] [37]

[38] [39] [40]

M. P. Merker et al. spin trap: a relatively superior trap for detecting superoxide anions and glutathiyl radicals. Free Radic. Biol. Med. 31:599 – 606; 2001. Merker, M. P.; Bongard, R. D.; Kettenhofen, N. J.; Okamoto, Y.; Dawson, C. A. Intracellular redox status affects transplasma membrane electron transport in pulmonary arterial endothelial cells. Am. J. Physiol. Lung Cell. Mol. Physiol. 282:L36 – L43; 2002. Liegeois, C.; Lermusieau, G.; Collin, S. Measuring antioxidant efficiency of wort, malt, and hops against the 2,2 V-azobis(2-amidinopropane) dihydrochloride-induced oxidation of an aqueous dispersion of linoleic acid. J. Agric. Food Chem. 48:1129 – 1134; 2000. Duling, B. R. Simulation of multiple isotropic spin-trap EPR spectra. J. Magn. Reson. 104:105 – 110; 1994. Chan, T. S.; Teng, S.; Wilson, J. X.; Galati, G.; Khan, S.; O’Brien, P. J. Coenzyme Q cytoprotective mechanisms for mitochondrial complex I cytopathies involves NAD(P)H: quinone oxidoreductase 1(NQO1). Free Radic. Res. 36:421 – 427; 2002. Penketh, P. G.; Hodnick, W. F.; Belcourt, M. F.; Shyam, K.; Sherman, D. H.; Sartorelli, A. C. Inhibition of DNA cross-linking by mitomycin C by peroxidase-mediated oxidation of mitomycin C hydroquinone. J. Biol. Chem. 276:34445 – 34452; 2001. Reszka, K. J.; McCormick, M. L.; Britigan, B. E. Oxidation of anthracycline anticancer agents by the peroxidase mimic microperoxidase 11 and hydrogen peroxide. Free Radic. Biol. Med. 35:78 – 93; 2003. Nunez, D. E.; Sanchez, F. A.; Garcia, C. F. A kinetic study of the one-electron oxidation of Trolox C by the hydroperoxidase activity of lipoxygenase. Biochim. Biophys. Acta 1335:127 – 134; 1997. Vanden Hoek, T. L.; Shao, Z.; Li, C.; Schumacker, P. T.; Becker, L. B. Mitochondrial electron transport can become a significant source of oxidative injury in cardiomyocytes. J. Mol. Cell. Cardiol. 29:2441 – 2450; 1997. Ohnishi, T.; Trumpower, B. L. Differential effects of antimycin on ubisemiquinone bound in different environments in isolated succinate – cytochrome c reductase complex. J. Biol. Chem. 255: 3278 – 3284; 1980. Forthoffer, N.; Gomez-Diaz, C.; Bello, R. I.; Buron, M. I.; Martin, S. F.; Rodriguez-Aguilera, J. C.; Navas, P.; Villalba, J. M. A novel plasma membrane quinone reductase and NAD(P)H:quinone oxidoreductase 1 are upregulated by serum withdrawal in human promyelocytic Hl-60 cells. J. Bioenerg. Biomembr. 34:209 – 219; 2002. Fato, R.; Estornell, E.; di Bernardo, S.; Pallotti, F.; Parenti, C. G.; Lenaz, G. Steady-state kinetics of the reduction of coenzyme Q analogs by complex I (NADH:ubiquinone oxidoreductase) in bovine heart mitochondria and submitochondrial particles. Biochemistry 35:2705 – 2716; 1996. Anusevicius, Z.; Sarlauskas, J.; Cenas, N. Two-electron reduction of quinones by rat liver NAD(P)H:quinone oxidoreductase: quantitative structure – activity relationships. Arch. Biochem. Biophys. 404:254 – 262; 2002. Moghaddas, S.; Hoppel, C. L.; Lesnefsky, E. J. Aging defect at the QO site of complex III augments oxyradical production in rat heart interfibrillar mitochondria. Arch. Biochem. Biophys. 414: 59 – 66; 2003. Powis, G.; Appel, P. L. Relationship of the single-electron reduction potential of quinones to their reduction by flavoproteins. Biochem. Pharmacol. 29:2567 – 2572; 1980. Goeptar, A. R.; Te Koppele, J. M.; van Maanen, J. M.; Zoetemelk, C. E.; Vermeulen, N. P. One-electron reductive bioactivation of 2,3,5,6-tetramethylbenzoquinone by cytochrome P450. Biochem. Pharmacol. 43:343 – 352; 1992. O’Brien, P. J. Molecular mechanisms of quinone cytotoxicity. Chem. Biol. Interact. 80:1 – 41; 1991. Floreani, M.; Carpenedo, F. Modifications of cardiac contractility by redox cycling alkylating and mixed redox cycling/alkylating quinones. J. Pharmacol. Exp. Ther. 256:243 – 248; 1991. La, M.; Rand, M. J. Effects of pyrogallol, hydroquinone and duroquinone on responses to nitrergic nerve stimulation and NO in the rat anococcygeus muscle. Br. J. Pharmacol. 126:342 – 348; 1999.

[41] Tanaka, T.; Okamura, T.; Handa, J.; Toda, N. Neurogenic vasodilation mediated by nitric oxide in porcine cerebral arteries. J. Cardiovasc. Pharmacol. 33:56 – 64; 1999. [42] Walles, S. A. Mechanisms of DNA damage induced in rat hepatocytes by quinones. Cancer Lett. 63:47 – 52; 1992. [43] Schmalix, W. A.; Lang, D.; Schneider, A.; Bocker, R.; Greim, H.; Doehmer, J. Stable expression and coexpression of human cytochrome P450 oxidoreductase and cytochrome P450 1A2 in V79 Chinese hamster cells: sensitivity to quinones and biotransformation of 7-alkoxyresorufins and triazines. Drug Metab. Dispos. 24:1314 – 1319; 1996. [44] Nemeikaite, A.; Cenas, N. The changes of prooxidant and antioxidant enzyme activities in bovine leukemia virus-transformed cells: their influence on quinone cytotoxicity. FEBS Lett. 326: 65 – 68; 1993. [45] Floreani, M.; Carpenedo, F. The catecholamine-mediated positive inotropic effect of simple quinones is related to superoxide anion generation. J. Pharmacol. Exp. Ther. 260:468 – 473; 1992. [46] Nakamura, M.; Hayashi, T. One- and two-electron reduction of quinones by rat liver subcellular fractions. J. Biochem. (Tokyo) 115:1141 – 1147; 1994. [47] Watanabe, N.; Forman, H. J. Autoxidation of extracellular hydroquinones is a causative event for the cytotoxicity of menadione and DMNQ in A549-S cells. Arch. Biochem. Biophys. 411:145 – 157; 2003. [48] Gardner, P. R. Superoxide production by the mycobacterial and pseudomonad quinoid pigments phthiocol and pyocyanine in human lung cells. Arch. Biochem. Biophys. 333:267 – 274; 1996. [49] Foster, C. E.; Bianchet, M. A.; Talalay, P.; Faig, M.; Amzel, L. M. Structures of mammalian cytosolic quinone reductases. [review]. Free Radic. Biol. Med. 29:241 – 245; 2000. [50] Butler, J.; Hoey, B. M. The one-electron reduction potential of several substrates can be related to their reduction rates by cytochrome P450 reductase. Biochim. Biophys. Acta 1161:73 – 78; 1993. [51] Beyer, R. E.; Segura-Aguilar, J.; di Bernardo, S.; Cavazzoni, M.; Fato, R.; Fiorentini, D.; Galli, M. C.; Setti, M.; Landi, L.; Lenaz, G. The role of DT-diaphorase in the maintenance of the reduced antioxidant form of coenzyme Q in membrane systems. Proc. Natl. Acad. Sci. USA 93:2528 – 2532; 1996. [52] Kagan, V. E.; Arroyo, A.; Tyurin, V. A.; Tyurina, Y. Y.; Villalba, J. M.; Navas, P. Plasma membrane NADH-coenzyme Q0 reductase generates semiquinone radicals and recycles vitamin E homologue in a superoxide-dependent reaction. FEBS Lett. 428:43 – 46; 1998. [53] Guo, Q.; Corbett, J. T.; Yue, G.; Fann, Y. C.; Qian, S. Y.; Tomer, K. B.; Mason, R. P. Electron spin resonance investigation of semiquinone radicals formed from the reaction of ubiquinone 0 with human oxyhemoglobin. J. Biol. Chem. 277:6104 – 6110; 2001. [54] Stocker, R.; Suarna, C. Extracellular reduction of ubiquinone-1 and -10 by human Hep G2 and blood cells. Biochim. Biophys. Acta 1158:15 – 22; 1993. [55] Corley, R. A.; English, J. C.; Hill, T. S.; Fiorica, L. A.; Morgott, D. A. Development of a physiologically based pharmacokinetic model for hydroquinone. Toxicol. Appl. Pharmacol. 165:163 – 174; 2000. [56] Bentinger, M.; Dallner, G.; Chojnacki, T.; Swiezewska, E. Distribution and breakdown of labeled coenzyme Q10 in rat. Free Radic. Biol Med. 34:563 – 575; 2003. [57] Yamashoji, S. Characterization of extracellular menadione-catalyzed H2O2 production by NIH/3T3 Cells. Biochem. Mol. Biol. Int. 44:555 – 563; 1998. [58] Thor, H.; Smith, M. T.; Hartzell, P.; Bellomo, G.; Jewell, S. A.; Orrenius, S. The metabolism of menadione (2-methyl-1,4-naphthoquinone) by isolated hepatocytes: a study of the implications of oxidative stress in intact cells. J. Biol. Chem. 257:12419 – 12425; 1982. [59] May, J. M.; Qu, Z. C.; Li, X. Ascorbic acid blunts oxidant stress due to menadione in endothelial cells. Arch. Biochem. Biophys. 411:136 – 144; 2003. [60] Nutter, L. M.; Ngo, E. O.; Fisher, G. R.; Gutierrez, P. L. DNA strand scission and free radical production in menadione-trea-

Endothelium and quinone redox status

[61]

[62] [63]

[64]

ted cells: correlation with cytotoxicity and role of NADPH quinone acceptor oxidoreductase. J. Biol. Chem. 267:2474 – 2479; 1992. Dikalov, S.; Landmesser, U.; Harrison, D. G. Geldanamycin leads to superoxide formation by enzymatic and non-enzymatic redox cycling: implications for studies of Hsp90 and endothelial cell nitric-oxide synthase. J. Biol. Chem. 277:25480 – 25485; 2002. Giulivi, C.; Cadenas, E. Extracellular activation of fluorinated aziridinylbenzoquinone in HT29 cells EPR studies. Chem. Biol. Interact. 113:191 – 204; 1998. Gant, T. W.; Rao, D. N.; Mason, R. P.; Cohen, G. M. Redox cycling and sulphydryl arylation: their relative importance in the mechanism of quinone cytotoxicity to isolated hepatocytes. Chem. Biol. Interact. 65:157 – 173; 1988. Brunmark, A.; Cadenas, E. Redox and addition chemistry of quinoid compounds and its biological implications. Free Radic. Biol. Med. 7:435 – 477; 1989.

103 ABBREVIATIONS

BMPO — 5-tert-butoxycarbonyl 5-methyl-1-pyrroline N-oxide BPAE — bovine pulmonary arterial endothelial cells DQ — duroquinone DQH2 — durohydroquinone DQS — durosemiquinone EPR — electron paramagnetic resonance HBSS — Hanks balanced salt solution HPLC — high performance liquid chromatography LDH — lactate dehydrogenase SOD — superoxide dismuatase

APPENDIX Under the assumption that oxidation of DQH2 by Fe(CN)6 3 in the medium is very rapid in comparison to the other DQH2 oxidation pathways, then

½FeðCNÞ6 3 ðtÞc½FeðCNÞ6 3 0  2½DQH2 ðtÞ ðA1Þ

for t > 0;

and Eq. (10) simplifies to





d½DQH2

1 V max1 ½DQ

k¯ 3 ¼ ðDQ þ dt V m K m1 þ ½DQ

Vm

  V max1 ½DQ 0 V max3 ½DQ 0 d½DQH2

1 c þ dt V m K m1 þ ½DQ 0 K m3 þ ½DQ 0   k¯ 3 : ðA5Þ 2V m ðk¯ 1 ½O2 þ k¯ 3 Þ Substituting Eq. (A5) in the derivative of Eq. (A1) with respect to time,

S Þ;  2

ðA2Þ where [Fe(CN)63]0 is the initial (t = 0) concentration of [Fe(CN)63] in cell medium. Assuming that for t > 0, a steady state would be reached in which [DQ ] was constant and small compared with [DQ], then

S

½DQ ðtÞc½DQ 0

where [DQ]0 is the initial (t = 0) concentration of DQ in cell medium. Substituting Eqs. (A3) and (A4) into Eq. (A2),

for t > 0;

ðA3Þ

  d½FeðCNÞ6 3 2 V max1 ½DQ 0 V max3 ½DQ 0 c þ dt Vm K m1 þ ½DQ 0 K m3 þ ½DQ 0   ¯k3 ¼ constant: 2ðk¯ 1 ½O2 þ k¯ 3 Þ ðA6Þ Thus, the model predicts zero-order kinetics for Fe(CN)63 when added to cell medium in the presence of DQ, which is consistent with the measured data (Figs. 3 and 5). For control conditions, where the two-electron pathway is dominant, Eq. (A6) simplifies to

and Eq. (11) simplifies to

ðDQS Þ

 2

  d½FeðCNÞ6 3 2 V max1 ½DQ 0 c ¼ constant: dt V m K m1 þ ½DQ 0

  V max3 ½DQ 0 1 ¼ ; K m3 þ ½DQ 0 2ðk¯ 1 ½O2 þ k¯ 3 Þ ðA4Þ

ðA7Þ