Kinetics of oxidation of tyrosine and dityrosine by myeloperoxidase compounds I and II. Implications for lipoprotein peroxidation studies.

The oxidation of lipoproteins is considered to play a key role in atherogenesis, and tyrosyl radicals have been implicated in the oxidation reaction. Tyrosyl radicals are generated in a system containing myeloperoxidase, H2O2, and tyrosine, but details of this enzyme-catalyzed reaction have not been explored. We have performed transient spectral and kinetic measurements to study the oxidation of tyrosine by the myeloperoxidase intermediates, compounds I and II, using both sequential mixing and single-mixing stopped-flow techniques. The one-electron reduction of compound I to compound II by tyrosine has a second order rate constant of (7.7 +/- 0.1) x 10(5) M-1 s-1. Compound II is then reduced by tyrosine to native enzyme with a second order rate constant of (1.57 +/- 0.06) x 10(4) M-1 s-1. Our study further revealed that, compared with horseradish peroxidase, thyroid peroxidase, and lactoperoxidase, myeloperoxidase is the most efficient catalyst of tyrosine oxidation at physiological pH. The second order rate constant for the myeloperoxidase compound I reaction with tyrosine is comparable with that of its compound I reaction with chloride: (4.7 +/- 0.1) x 10(6) M-1 s-1. Thus, although chloride is considered the major myeloperoxidase substrate, tyrosine is able to compete effectively for compound I. Steady state inhibition studies demonstrate that chloride binds very weakly to the tyrosine binding site of the enzyme. Coupling of tyrosyl radicals yields dityrosine, a highly fluorescent stable compound that had been identified as a possible marker for lipoprotein oxidation. We present spectral and kinetic data showing that dityrosine is further oxidized by both myeloperoxidase compounds I and II. The second order rate constants we determined for dityrosine oxidation are (1.12 +/- 0.01) x 10(5) M-1 s-1 for compound I and (7.5 +/- 0.3) x 10(2) M-1 s-1 for compound II. Therefore, caution must be exercised when using dityrosine as a quantitative index of lipoprotein oxidation, particularly in the presence of myeloperoxidase and H2O2.

A major risk factor for coronary artery disease is an elevated level of low density lipoproteins (LDL), 1 the major carrier of blood cholesterol (1). Thus, the reduction of LDL levels has been a prominent preventive measure against atherosclerosis. However, evidence has accumulated indicating that oxidized LDL rather than native LDL triggers the pathological events leading to atherosclerosis (1)(2)(3)(4). Foam cells or lipid-laden macrophages, derived from circulating monocytes, are the earliest cellular components of atherosclerotic lesions (5). It has been found that normal macrophages in culture cannot be converted to foam cells by simple incubation with even very high concentrations of native LDL. It is postulated that circulating LDL first undergo oxidative modification and then the oxidized LDL convert normal macrophages into foam cells (6,7).
Myeloperoxidase, a heme protein abundant in phagocytes (8), is a potential physiological catalyst for lipoprotein oxidation (9,10). Recent studies demonstrate links between myeloperoxidase and oxidative damage to proteins and lipids (11,12). The identification of myeloperoxidase in human atherosclerotic lesions (13) is support for the hypothesis that myeloperoxidase is indeed involved in lipoprotein oxidation.
While myeloperoxidase follows the normal peroxidase cycle (14,15), MPO  where a single two-electron oxidation of native enzyme (MPO) to compound I (MPO-I) is followed by two successive one-electron reductions back to native enzyme via compound II (MPO-II), it shares an ability with eosinophil peroxidase (16) among the mammalian peroxidases in utilizing chloride to produce hypochlorous acid (17,18), HOCl is a potent oxidizing agent that plays a cytotoxic role against invading bacteria, viruses, and tumor cells (8). However, it is also known to injure normal tissues by bleaching heme groups and oxidatively destroying electron transport chains (19) as well as chlorinating amino acids and unsaturated lipids (10,20,21). HOCl secreted by activated neutrophils can also cause oxidation of lipoproteins in vivo (22). Moreover, myeloperoxidase has been found to convert cholesterol to chlorohydrins and epoxides by a reaction involving HOCl (11).
Another potential lipoprotein oxidative pathway involving myeloperoxidase implicates the tyrosyl radical produced via one-electron oxidation of tyrosine by MPO-I (12,23). When two tyrosyl radicals combine the major product is o,oЈ-dityrosine, an intensely fluorescent compound (24,25). Fluorescence due to dityrosine has been observed in the reaction of tyrosine with linoleic acid 13-monohydroperoxide in the presence of methemoglobin (26). Dityrosine has also been detected in high density lipoproteins (HDL) treated with horseradish peroxidase, hydrogen peroxide, and tyrosine (27).
The concentration of free tyrosine in human blood plasma is 55 M (28) and ranges from 22 to 83 M in venous blood (29). Since dityrosine formation has been observed even in the presence of physiological concentrations of chloride (12,30), the major MPO substrate, it has been suggested that tyrosine can be an effective reducing substrate for MPO. While the literature abounds with evidence for oxidative tyrosylation of lipoproteins (12,23,27,30), limited if any details are reported on the myeloperoxidase-catalyzed one-electron oxidation of tyrosine. In this paper we report spectral and transient kinetic data for the oxidation of tyrosine by the MPO/H 2 O 2 system. Rate constant determination for MPO-I reactions were made possible by employing sequential mixing stopped-flow techniques. We also report rate constants for tyrosine oxidation by MPO-II. The contribution of this reaction to tyrosyl-mediated lipid peroxidation has never received attention to the best of our knowledge. Moreover, the retention of the phenolic groups in the tyrosine oxidation product, dityrosine, raises the possibility of further oxidation. Herein we also report rate constants for dityrosine oxidation by both MPO-I and MPO-II. These data will be useful in evaluating the stability of the end product dityrosine, which has been proposed as a possible marker for lipid peroxidation (30) or as an index of oxidative damage to proteins in vivo and in vitro (31,32). Since chloride is considered the major substrate of myeloperoxidase in vivo, we also investigated the steady state rates of tyrosine oxidation in the presence and absence of chloride. By use of a sequential-mixing stopped-flow instrument we have measured the rate constant for MPO-I reaction with chloride previously unreported in the literature. These results not only contribute quantitative support to the accumulating evidence on the role of tyrosyl radicals in lipoprotein peroxidation but also present differences in tyrosine oxidation by MPO compared with other peroxidases.

EXPERIMENTAL PROCEDURES
Materials-Bovine spleen myeloperoxidase was isolated and purified using a combination of modified procedures (33)(34)(35)(36). The enzyme preparation used in this study exhibited an RZ (A 429 /A 280 ) of 0.83. The MPO concentration was determined spectrophotometrically using an extinction coefficient of 178 mM Ϫ1 cm Ϫ1 at 429 nm (37).
Diluted hydrogen peroxide, obtained as a 30% solution from BDH Chemicals, was standardized using the horseradish peroxidase-catalyzed oxidation of iodide to triiodide (38). Concentrations were confirmed using absorbance measurements at 240 nm where the extinction (39). Ultrapure L-tyrosine was obtained from Sigma. KCl (Aldrich) and the chemicals used for the buffers (Fisher) were used without further purification. Aqueous solutions were prepared using water purified through the Milli-Q system (Millipore Corp.), and concentrations of solutes were determined by weight.
Dityrosine was prepared using a combination of published procedures (24,40,41). 1.0 gram of L-tyrosine was dissolved in 910 ml of water, and then 50 ml of 0.1% H 2 O 2 and 40 mg of horseradish peroxidase (Type VI-A, Sigma) were added. The pH was adjusted to 9.2 using 6 M NaOH, and the mixture was incubated in a thermostated bath for 17 h at 37°C. At the end of the reaction the pH was adjusted to 6.0 with concentrated HCl. The mixture was concentrated almost to dryness in a rotatory evaporator under vacuum at 40°C. Water was added to the concentrate to a volume of 60 ml. The mixture was treated with 2 g of Darco G-60 activated charcoal (Aldrich) and left standing overnight. Centrifugation at 10,000 rpm for 15 min using the JA-20 rotor of Beckman model J2-21 preparative centrifuge yielded a clear yellowish brown supernatant. This was then applied to a precycled P-11 cellulose phosphate cationic exchanger (Whatman) column (1.5 ϫ 35 cm), which had been previously equilibrated with 0.2 M acetic acid. Elution of the column was performed with 0.2 M acetic acid containing 0.5 M NaCl at a flow rate of 15 ml/h. Fractions of 3 ml were collected and evaluated spectrophotometrically at 280 and 310 nm. Samples of the major peak fractions were transferred on thin layer silica gel plates (Whatman). After developing the plates in n-butyl alcohol/acetic acid/water (4:1:1, v/v) the fractions were pooled that were positive for ninhydrin and with R f value (the ratio of the distance traveled by the band to the distance traveled by the mobile phase) of about 0.15. The solution was applied to a Dowex 50 ϫ 8 (Terochem) column previously soaked in 0.1 M HCl. The column was extensively washed with water to remove all NaCl and acetic acid. The dityrosine was subsequently eluted with 2 M ammonia. Eluates were evaluated spectrophotometrically as before. Samples of major peak fractions were applied on thin layer silica gel plates, and the fractions with R f values of about 0.12 were pooled. The solvent was removed in a rotatory evaporator under vacuum. The concentrate of dityrosine was characterized by thin layer chromatography, ultraviolet spectroscopy and fluorescence measurements (42). Final concentrations were determined by measuring absorbance at 315 nm using an extinction coefficient of 5.2 mM Ϫ1 cm Ϫ1 at pH 7.5 (43).
Methods-Routine absorbance measurements were made on a Beckman DU-650 spectrophotometer equipped with thermally jacketed 1-cm cuvette holders. Rapid spectral scans and kinetic measurements were performed using the SX.17 MV microvolume stopped-flow spectrofluorimeter (Applied Photophysics). Time-dependent spectra from single wavelength shots were reconstructed using the GLint application software. A 1-cm path length was used for absorbance measurements, and a 2-mm path length was used for fluorescence measurements. All experiments were carried out at 20.5 Ϯ 0.5°C, ionic strength of 0.1 M, and pH 7.4 (phosphate buffer) unless otherwise specified. All pH measurements were made using a Fisher Accumet model 25 digital pH meter.
The kinetics of the reactions of MPO-I with tyrosine and dityrosine were carried out using the sequential mixing mode of the stopped-flow apparatus. 1.0 M of myeloperoxidase was premixed with 20 M H 2 O 2 in 0.1 M phosphate buffer. After a delay time of 20 ms the formed MPO-I was allowed to react with varying concentrations of L-tyrosine or dityrosine, the final concentrations of which were at least 10-fold in excess of the enzyme intermediate. The time course of the reaction was followed by monitoring the absorbance changes at 456 nm accompanying the formation of MPO-II. This wavelength is also the isosbestic point between native enzyme and MPO-I (36).
The reaction of MPO-I with chloride was investigated at 429 nm, the isosbestic point between MPO-I and MPO-II, which also corresponds to the maximum change in absorbance as MPO-I undergoes a two-electron reduction back to the native enzyme. Pseudo-first order rate constants, k obs , were determined using the single exponential curve-fit equation of the Applied Photophysics software. Five or six determinations of rate constants were performed for each reducing substrate concentration, and the mean values were plotted against the substrate concentration. The apparent second order rate constants for the MPO-I reactions were calculated from the slopes using linear least squares regression analysis (Enzfitter, Elsevier-Biosoft).
MPO-II reactions with L-tyrosine and dityrosine were carried out using the single-mixing mode of the stopped-flow apparatus. Prior to the kinetic experiments the stability of preformed MPO-II was studied by wavelength scanning using the Beckman spectrophotometer. MPO-II was prepared by adding a 50-fold excess of H 2 O 2 to native enzyme and then allowing this enzyme intermediate to react with tyrosine and dityrosine under pseudo-first order conditions. Kinetic measurements were made at 456 nm where the maximum change in absorbance is observed as MPO-II is converted back to native enzyme. Pseudo-first order rate constants were determined from seven or more exponential traces, and second order rate constants were calculated as before.
Steady state experiments on the oxidation of tyrosine by the MPO/ H 2 O 2 system were conducted by following the initial rate of formation of dityrosine. The reaction was monitored by measuring the increase in fluorescence with time using wavelengths of 325 and 405 nm for excitation and emission. In a typical experiment one syringe contained MPO and tyrosine in phosphate buffer while the other contained H 2 O 2 .
To determine the effect of chloride, known concentrations of KCl solution were added to the syringe containing H 2 O 2 . The final concentrations after mixing were 25 nM MPO, 50 M H 2 O 2 , 0.2-1.0 mM tyrosine, and 0 -0.125 M KCl. The initial rates were determined from the slopes of the first linear portions of the traces obtained after mixing all the components. The dead time of the instrument was ϳ1.5 ms. Usually six to eight traces were recorded, and the mean values of the initial rate of increase in fluorescence with time were calculated. A calibration curve of fluorescence signal (volts) against dityrosine concentration is linear up to 20 M dityrosine. By interpolation, initial rates in fluorescence units were converted to rates in terms of dityrosine concentration. Double reciprocal plots as well as Dixon plots were constructed to determine the type of inhibition chloride exerts on tyrosine oxidation (44).

RESULTS
Preliminary wavelength scans of myeloperoxidase in phosphate buffer in the absence and presence of tyrosine or dityrosine revealed no interference in the wavelengths to be used for transient kinetic measurements (spectra not shown). Kinetic scans of the reaction between native MPO and H 2 O 2 in the presence of tyrosine are shown in Fig. 1. The first scan is that of native enzyme, and the second scan is that of MPO-I. The succeeding scans demonstrate the rapid reduction of MPO-I to MPO-II. MPO-I formation takes place within 10 ms after mixing, followed by its conversion to MPO-II, which is completed in 100 ms. Full formation of MPO-I is achieved only in the presence of a 20-fold excess of H 2 O 2 (36). However, since MPO-I is also spontaneously reduced to MPO-II in the presence of excess H 2 O 2 it was necessary to determine the rate of reduction of MPO-I by tyrosine relative to its rate of spontaneous decay. The instability of MPO-I necessitated the use of the sequential mixing stopped-flow apparatus. The premixing of MPO and H 2 O 2 led to MPO-I formation within 20 ms, and before the MPO-I decayed it allowed the measurement of the rate of the subsequent reaction of MPO-I with a reducing substrate.
The time courses for the reduction of MPO-I to MPO-II may be followed by monitoring the disappearance of MPO-I at 442 nm, the isosbestic point between MPO and MPO-II, or the formation of MPO-II at 456 nm, the isosbestic point between MPO and MPO-I. Rate constants obtained in both cases are the same within experimental uncertainty (36). We chose to perform kinetic measurements for MPO-I reduction by tyrosine at 456 nm because the absorbance change accompanying MPO-II formation at this wavelength is larger and leads to smaller experimental errors. A typical kinetic trace displaying single exponential character is shown in the inset to Fig. 2. The apparent second order rate constant was obtained from the slope of the plot of pseudo-first order rate constants, k obs , against tyrosine concentration (Fig. 2): (7.7 Ϯ 0.1) ϫ 10 5 M Ϫ1 s Ϫ1 . A value of (6.3 Ϯ 0.2) ϫ 10 5 M Ϫ1 s Ϫ1 was obtained using the single-mixing mode of the apparatus. However, the intercept of the plot obtained using the single-mixing mode is bigger, 9.5 Ϯ 1.8 s Ϫ1 , compared with 2.2 Ϯ 1.2 s Ϫ1 (Fig. 2), obtained using the sequential-mixing mode. The value of the intercept indicates the rate of the spontaneous reduction of MPO-I by excess H 2 O 2 in the absence of tyrosine. Thus, the use of sequential mixing stopped-flow minimized the interference of MPO-I reduction to MPO-II by excess H 2 O 2 .
The rate constant for MPO-I reduction by tyrosine at pH 7.4 was higher than the rate at pH 4.5, the pH inside the phagocyte (45). We performed pseudo-first order rate measurements at single tyrosine concentrations from pH 3.0 to 8.8, and the results are presented in Fig. 3.
While MPO-I is thermodynamically competent to form tyrosyl radicals via its one-electron oxidation of tyrosine (17), the capability of MPO-II to generate tyrosyl radicals was unknown (12). MPO-II is more stable than MPO-I (36,46,47). A good preparation of MPO-II is obtained by adding a 50-fold excess of H 2 O 2 to pure native enzyme. The MPO-II spectra remained basically unchanged after 10 min (Fig. 4). Upon addition of tyrosine, MPO-II is rapidly reduced to native enzyme. This one-electron enzyme reduction must be accompanied by a oneelectron oxidation of the reducing substrate. These results prove that MPO-II is also capable of oxidizing tyrosine and generating tyrosyl radicals.
Because of the relative stability of MPO-II we were able to conduct rate measurements of the reaction of MPO-II with tyrosine using the single-mixing mode of the stopped-flow apparatus. The monophasic exponential decrease in absorbance at 456 nm with time as MPO-II is converted back to native enzyme yielded k obs values that were plotted against tyrosine concentration as before (data not shown). The plot was linear up to 250 M tyrosine, and the intercept is zero within the standard deviations of the measurements. The second order rate constant for the reaction of MPO-II with tyrosine obtained from the slope of the plot was (1.57 Ϯ 0.06) ϫ 10 4 M Ϫ1 s Ϫ1 .
Dityrosine has been detected in the MPO/H 2 O 2 system of human neutrophils and macrophages as a product of tyrosyl radical coupling (30). Aside from the dimer, other polymeric products have been identified in sytems employing horseradish peroxidase (25) and ovoperoxidase (48). It was of interest to determine whether MPO would further catalyze the oxidation of dityrosine since the phenolic groups are retained in the dimer.
Dityrosine was prepared enzymatically, purified, and characterized (42). For the kinetic measurements it is important to work with authentic dityrosine freed of tyrosine impurities. Dityrosine is easily monitored by its characteristic fluorescence at 400 nm (49). It also shows maximum absorbance at 285 nm in acid solution and at 315 in alkaline solution (26). Dityrosine yields a single ninhydrin-positive spot on silica gel with R f of 0.12 when using n-butyl alcohol/acetic acid/water (4:1:1 v/v) as a solvent system (42,50). All of these criteria were satisfied by the dityrosine used in this work.
The addition of dityrosine to preformed MPO-II resulted in conversion of MPO-II to native enzyme, although not as fast compared with the reaction of tyrosine. A shoulder at 456 nm in the wavelength scans (Fig. 5, inset) indicates that MPO-II reduction is not complete within the time period of the scan. Similar rate measurements under pseudo-first order conditions were conducted for the reactions of MPO-I and MPO-II with dityrosine. The second order rate constant determined from the slope of the plot of k obs against dityrosine concentration for the MPO-II reaction is (7.5 Ϯ 0.3) ϫ 10 2 M Ϫ1 s Ϫ1 (Fig. 5). The second order rate constant for the MPO-I reaction with dityrosine was more than 2 orders of magnitude larger: (1.12 Ϯ 0.01) ϫ 10 5 M Ϫ1 s Ϫ1 .
The major reducing substrate for MPO is considered to be chloride, the oxidation product of which is hypochlorous acid (8). It has been suggested that chloride inhibits the initial rate of oxidation of tyrosine by myeloperoxidase (30). It is relevant to evaluate the relative rates of tyrosine and chloride oxidation by MPO-I. By using sequential mixing stopped-flow measure-ments we determined the second order rate constant for MPO-I oxidation of chloride to be (4.7 Ϯ 0.1) ϫ 10 6 M Ϫ1 s Ϫ1 . Unlike previous plots, which had nearly zero intercepts (Figs. 2 and 5, for example), the plot of k obs versus chloride concentration yielded an intercept of (31 Ϯ 4) s Ϫ1 (data not shown).
We were also interested in the type of inhibition chloride exerts on tyrosine oxidation. This was investigated under steady state conditions by monitoring the rate of formation of the dityrosine oxidation product in a system containing MPO, H 2 O 2 , tyrosine, and chloride. The rate of dityrosine formation may be followed by monitoring absorbance changes at 315 nm (43) or fluorescence changes using 325 and 405 nm as excitation and emission wavelengths. The fluorescence signal is larger, so we chose it over absorbance to measure initial rates of dityrosine formation. Fig. 6 shows that tyrosine does not contribute to the fluorescence reading. The concentration of dityrosine in the reaction mixture was interpolated from a calibration curve of purified dityrosine (Fig. 6, inset).
The initial rate of tyrosine oxidation was measured as a function of tyrosine concentration, and the results are shown in Fig. 7. Obviously, the plot is not hyperbolic and therefore does not yield corresponding Michaelis-Menten parameters. Double reciprocal plots of initial rates against tyrosine concentration increase in slope as the concentration of chloride increases (Fig.  8). A secondary plot of the slopes from the double reciprocal plots against chloride concentration (Fig. 8, inset) yields an inhibitor constant K i equal to 0.16 M. Values of K i are usually obtained accurately by using Dixon plots (51), i.e. plots of the reciprocal of initial rate against inhibitor concentration at various fixed concentrations of substrate. The family of Dixon plots in Fig. 9 suggests either competitive or linear mixed-type inhibition (44), and from the point of intersection we obtained the inhibition constant of chloride, K i ϭ 0.18 Ϯ 0.02 M; and the maximum velocity for tyrosine oxidation, V max ϭ 1.

DISCUSSION
The peroxidase-catalyzed oxidation of tyrosine has recently attracted renewed interest due to its involvement in lipid and protein oxidation (12,27,31). The oxidation of tyrosine has also been implicated in the biosynthesis of thyroxine and melanin (43) and in the cross-linking of structural proteins via the dityrosine linkage (for a review see Ref. 42). Previous investigations of the kinetics of tyrosine oxidation have been performed using lactoperoxidase (43,52), horseradish peroxidase (53,54), and thyroid peroxidase (52,55). Our present work provides the first report of the rate constants for the oxidation of tyrosine as catalyzed by myeloperoxidase. These results are significant in lieu of the findings that MPO has been identified as a physiological catalyst for lipid peroxidation in low density lipoproteins via tyrosyl radicals (12,23).
While all hemoprotein peroxidases are generally considered to catalyze oxidation reactions in a similar manner, Table I shows varying values in the rate constants for the oxidation of tyrosine by several peroxidases under similar pH and temperature conditions. This reflects differences in the effectiveness of various peroxidases in catalyzing the oxidation of tyrosine. Horseradish peroxidase is the least effective catalyst for tyrosine oxidation. Among the three mammalian peroxidases, thyroid peroxidase has the lowest rate constant for compound I reduction by tyrosine. This is to be expected because the primary reaction catalyzed by thyroid peroxidase is iodination of the tyrosyl residues in thyroglobulin; therefore, tyrosine oxidation may not be as important. Moreover, thyroid peroxidase has a unique property of catalyzing the two-electron oxidation of tyrosine (55), which differentiates it from lactoperoxidase and myeloperoxidase, which oxidize tyrosine via two successive one-electron processes. The rapid spectral scans of the MPO/ H 2 O 2 system in the presence of tyrosine (Fig. 1) clearly show   FIG. 6. Fluorescence spectra of dityrosine. The excitation wavelength was set at 325 nm. Tyrosine does not fluoresce when exposed to 325-nm light. The inset shows the calibration curve used for calculating dityrosine concentration from fluorescence readings.   that MPO-I undergoes a one-electron reduction to MPO-II followed by the slower one-electron reduction of MPO-II to native enzyme (Fig. 4). The rate constant for lactoperoxidase compound-I reaction with tyrosine could only be estimated by previous workers (52) due to the inherent instability of its compound I. We were able to overcome this problem in our transient kinetic measurements through the use of a sequential mixing stopped-flow apparatus. Our data establish that myeloperoxidase is the most effective among the mammalian peroxidases in catalyzing both the one-electron reductions of its compounds I and II by tyrosine. Our results also show that tyrosine could contribute to the turnover of MPO-II to native enzyme in the absence of other reducing substrates.
Another striking difference displayed by these peroxidases in their oxidation of tyrosine is the pH at which the maximum rate is observed for compound I reduction. For horseradish peroxidase, the maximum rate is at pH 9.6 (53), while for lactoperoxidase it is at pH 8.2 (43). Fig. 3 indicates that for MPO, the maximum rate is achieved at the physiological pH of 7.4. In a related study of the pH dependence, the formation of dityrosine, the stable primary product of the myeloperoxidasecatalyzed oxidation of tyrosine, was found to be highest at pH 7.5-8.0 (30). Apparently, oxidation of tyrosine requires deprotonation of its phenolic hydroxyl group (56). Thus, inside the acidic environment of the phagosome (45) the predominant reaction is not likely tyrosine oxidation but rather chloride oxidation by MPO-I to form HOCl. Moreover, it has been demonstrated that only at acidic pH is there direct contact between chloride and ferryl oxygen in MPO-I resulting in a maximum rate of HOCl formation (57).
MPO-I and MPO-II are each able to oxidize tyrosine in a one-electron reaction to give tyrosyl radicals. Long-lived tyrosyl radicals have also been produced by chemical oxidation of tyrosine using ferricyanide and characterized by electron spin resonance (58). However, the readily detectable product of tyrosine oxidation is o,oЈ-dityrosine formed by coupling of phenoxy radicals (25). Dityrosine is intensely fluorescent (Fig. 6), and in proteins it has been found to be resistant to proteolysis (31), to borohydride treatment (26), and to acid hydrolysis (30). Because of its apparent stability dityrosine has been proposed as a chemical marker of cumulative exposure of proteins to metal-catalyzed oxidation in vitro and in vivo (32), an index of organismal oxidative stress (31), and an indication of targets where phagocytes inflict oxidative damage in vivo (30). However, it is also known that extensive action of peroxidases on tyrosine results in the formation of other oxidation products such as trityrosine and other polymers, some of which are also fluorescent (25,30,43,48). This is probably the reason why the steady state rates of dityrosine formation measured by the increase in fluorescence did not reach saturation but continued to increase with time (Fig. 7).
Since the phenoxy groups are retained in dityrosine it is likely that dityrosine can be further oxidized by the MPO/H 2 O 2 system. The inset to Fig. 5 shows that dityrosine can reduce MPO-II to native enzyme, although the rate would not seem physiologically relevant. On the other hand the rate constant for MPO-I reaction with dityrosine yields a value, (1.12 Ϯ 0.01) ϫ 10 5 M Ϫ1 s Ϫ1 , which is not significantly lower than the reaction of MPO-I with tyrosine. The lower rate constant for the more bulky dityrosine substrate is consistent with the finding that aromatic substrate molecules bind at the distal heme pocket of myeloperoxidase (59) and that this site exhibits considerable steric hindrance (17). The fast oxidation of dityrosine by MPO-I suggests that dityrosine may not be a very stable end product and that it should be used with caution as a marker for oxidative damage, particularly in the presence of the MPO/ H 2 O 2 system.
Since the major physiological substrate of MPO is considered to be chloride, we also evaluated the relative rates of tyrosine and chloride oxidation by MPO-I. There have been difficulties in obtaining the rate constant for the two-electron oxidation of chloride by MPO-I. Full formation of MPO-I is achieved only in the presence of at least a 20-fold excess of H 2 O 2 over MPO (30). But with excess H 2 O 2 , MPO-I is reduced at a very fast rate to MPO-II, making it difficult to measure the rate constant for the MPO-I reaction with chloride. We were able to overcome this problem by using the sequential mixing mode of the stoppedflow apparatus. The second order rate constant for chloride peroxidation by MPO-I, (4.7 Ϯ 0.1) ϫ 10 6 M Ϫ1 s Ϫ1 , is about an order of magnitude higher than the rate constant for tyrosine oxidation by MPO-I and about the same value as the rate constant for the formation of an MPO-chlorinating intermediate (2.8 Ϯ 1.2) ϫ 10 6 M Ϫ1 s Ϫ1 (60). However, unlike the other plots of k obs against substrate concentration, which had zero intercepts within experimental error (Figs. 2 and 5, for example), the plot for the chloride reaction with MPO-I yielded a considerable y intercept (31 Ϯ 4) s Ϫ1 . This suggests a reversible reaction. In fact, the oxidation product of chloride, HOCl, can undergo a rapid reaction with native MPO to give MPO-I, which is ostensibly the reverse of the chloride peroxidation reaction (17,18).
For the chloride inhibition studies we constructed both double reciprocal and Dixon plots, which are used to identify the type of inhibition and to determine the inhibition constant, K i (51). The two types of plots only establish that the inhibition is neither noncompetitive nor uncompetitive. The data are characteristic of either purely competitive or mixed-type inhibition (44). The replot of the slopes of the Dixon plots against reciprocal of substrate (tyrosine) concentration did not umambiguously predict which type of inhibition is occurring. The Cornish-Bowden plot (61) (graphs not shown) did not clarify the kind of inhibition either. Indeed it may be difficult to characterize the type of inhibition in such a complex system where the enzyme is in contact with three substrates: H 2 O 2 , tyrosine, and chloride. A simple interpretation of plots used for inhibition studies is not possible for multisubstrate systems (61), and especially where the products (HOCl and dityrosine) are also substrates. H 2 O 2 and chloride have been shown to be competitive inhibitors of each other (62)(63)(64). The substrate inhibition patterns of these studies suggest the existence of two binding sites for chloride; one is a substrate binding site and the other an inhibitor binding site. The binding of chloride to its substrate binding site results in the production of HOCl, whereas its binding to the inhibitor binding site leads to competitive inhibition of H 2 O 2 binding to native MPO. Inhibition by chloride almost certainly involves coordination to the heme because MPO-I has a ferryl oxygen occupying the sixth coordination position and possesses no available chloride binding sites. The catalytic site for chloride peroxidation is most likely located within the distal heme cavity (57) or at the heme periphery (17,65). Recently, EPR spectroscopy and model building revealed that the hydrophobic character of the entrance to the distal cavity of MPO allows favorable interaction with aromatic molecules (59) such as tyrosine. If indeed the chloride substrate binding site is in the distal heme pocket, then it would compete with tyrosine. However, the large value of K i we obtained in our steady state studies suggests that binding of chloride is very weak, presumably because of the hydrophobic nature of the pocket. That would explain why tyrosine oxidation takes place at considerable rates even in the presence of high concentrations of chloride. These results are consistent with the findings that tyrosine oxidation occurs even in the presence of physiological concentrations of chloride (12,30) and that LDL oxidation by tyrosyl radicals is largely unaffected by as high as 0.1 M chloride present in the plasma (23).
In conclusion, both MPO-I and MPO-II are able to react with tyrosine via one-electron oxidations and therefore generate tyrosyl radicals. The relatively fast rate of the MPO-I reaction with tyrosine substantiates the conjecture that tyrosine oxidation by MPO-I takes place in vivo and may be physiologically relevant. Tyrosine also competes very effectively with the major myeloperoxidase substrate chloride. That tyrosine can also reduce MPO-II to native enzyme ensures turnover of the enzyme even in the absence of other reducing substrates. The tyrosyl radical formed in both reactions is a diffusible catalyst that can convey oxidizing potential away from the active site of the heme enzyme. A protein tyrosyl radical has been implicated in cross-linking of proteins (12), in initiating lipid peroxidation in LDL (23) and modifying HDL to protect the arterial wall against cholesterol accumulation (27). The readily detectable product of tyrosine oxidation is dityrosine formed by the coupling of tyrosyl radicals. Our study revealed that since dityrosine is further oxidized at a very fast rate by MPO-I caution must be exercised when using it as a quantitative index of lipoprotein modification. It would appear to measure a lower limit of lipoprotein modification.