ARCHIVES
OF BIOCHEMISTRY
Vol. 299, No. 1, November
AND
BIOPHYSICS
15, pp. 47-56,
1992
Electron Spin Resonance Study of Peroxidase Activity and Kinetics’ Katherine
L. Moore,
Division
of Energy
Received
April
Mario
M. Moronne,
and Environment,
22, 1992. and in revised
Lawrence
form
July
and Rolf J. Mehlhorn’ Berkeley
Laboratory,
Horseradish peroxidase (HRP)3 and lactoperoxidase (LPO) are typical of peroxidase enzymes whose catalytic ’ The U.S. Government’s right to retain a nonexclusive royalty-free license in and to the copyright covering this paper, for governmental purposes, is acknowledged. ’ To whom correspondence should be addressed at Division of Energy and Environment, M/S 7O-193A, Lawrence Berkeley Laboratory, 1 Cyclotron Road, Berkeley, CA 94720. 3 Abbreviations used: HRP, horseradish peroxidase; LPO, lactoperoxidase; ABTS, 2,2’-azinobis(3-ethylbenzthiazoline-6-sulfonic acid; TOLH, 1,4-dihydroxy-2,2,6,6-tetramethylpiperadine; Tempol, 2,2,6,6tetramethyl-l-piperidinoxy-4-01; PBS, phosphate-buffered saline; DTPA, diethylene triaminepentaacetic acid; TPO, thyroid peroxidase.
$5.00
Copyright 8 1992 by Academic Press, Inc. All rights of reproduction in any form reserved.
California
94720
22, 1992
An electron spin resonance (ESR) assay has been developed for peroxidase activity. The assay measures the formation of the paramagnetic nitroxide Tempo1 from the oxidation of its hydroxylamine derivative (TOLH) by short-lived radicals produced by peroxidase cycle intermediates, Compounds I and II. Using phenol as a peroxidase electron donor, the ESR approach is suitable for measurements of peroxidase activity (aO.003 U/ml) and micromolar quantities of HzOz in sample sizes as small as 2 ~1. In addition, the ESR method can be used to continuously monitor activity in cell suspensions and other media that are susceptible to optical artifacts. The high membrane permeability of TOLH also makes it possible to estimate peroxidase activity in membrane-enclosed compartments, provided that TOLH oxidation rates can be stimulated with exogenous peroxidase reductants, e.g., phenol. Analysis of TOLH oxidation rates under conditions of low electron donor concentrations and high concentrations of HzOz also shows clear indications of substrate-dependent inhibition and increased catalatic activity. Computer simulations indicate that the results obtained are consistent with the peroxidase reaction scheme proposed by Kohler et al. (1988, Arch. Biochem. Biophys. 264, 438-449) modified to correct for a nitroxide dependent stimulation of peroxidase catalytic activity. ‘tt 1992 Academic Press, Inc.
oow9861/92
Berkeley,
cycle involves a number of distinct intermediates with differing oxidation states of the active iron center (for reviews see (1) and (2)). Native enzyme (ferriperoxidaseFe3+) reacts with HzOz to form Compound I (Fe5+), which may then return to the native enzyme by either a single two-electron reduction or by two one-electron steps. The source of donor electrons determines the path and rate of Compound I conversion back to native enzyme. Compound I is rapidly reduced by iodide in a single two-electron process (3). At a much slower rate, H,Oz may also serve as a two-electron donor largely accounting for the modest catalatic activity of peroxidases (3, 4). In the absence of an exogenous electron donor, Compound I can return to native enzyme in two relatively slow oneelectron reactions with reducing groups contained within the enzyme. For HRP, this process can proceed through 10 or more cycles (5). Exogenous one-electron donors (e.g., phenol) greatly increase enzyme turnover by facilitating transformation of Compound I to Compound II and Compound II back to native enzyme. In fact, a variety of inorganic and organic electron donors, including sulfur compounds and halogenated and nonhalogenated phenols, are suitable substrates for these peroxidase intermediates (2, 6). Various studies (3, 4, 7) have shown that H,OZ, in addition to serving as a substrate, can inhibit the catalytic cycle by reacting with Compound II to form Compound III, which is outside the peroxidatic loop (see Fig. 1). This has been proposed to occur by two different pathways, one of which depends on the formation of superoxide (4, 8). Subsequent to formation, Compound III can release oxygen to yield the ferrous enzyme, which further reacts with Hz02 to regenerate Compound II (3,9). Alternatively, Compound III may irreversibly decompose, resulting in a permanent loss of peroxidase activity (3). In the presence of appropriate electron donors such as 2,2’-azinobis(3ethylbenzthiazoline-6-sulfonic acid) (ABTS), the formation of Compound III can be reduced (7), thereby decreasing enzyme inactivation when exposed to high con47
48
MOORE,
MORONNE,
centrations of HzOz. This last feature is important in the optimal design of systems to monitor peroxidase activity in that conditions that conserve the enzyme can be chosen. In this paper, we present the results of an electron spin resonance (ESR) study of the peroxidase reaction cycle. In particular, the reaction cycle is evaluated using an ESR assay for the one-electron oxidation products of the peroxidase . HzOz complex using the reduced hydroxylamine derivative of the spin probe Tempo1 (TOLH). The approach taken depends on the capacity of Compounds I and II to rapidly oxidize phenolic substrates, producing radical intermediates which in turn oxidize the reduced nitroxide to its radical form. In this process, the phenol is regenerated and the accumulation of the nitroxide monitored by ESR spectrometry. With this approach, peroxidase activity > 0.003 U/ml is readily measured. Although accepted techniques for evaluation of peroxidase activity already exist (e.g., the guaiacol assay, Ref. (lo)), previous methods have largely depended on optical measurements that are unsuitable for many biological preparations such as turbid cell or microsomal suspensions. ESR is essentially free of optical artifacts, and its high sensitivity and small sample requirements (2-20 ~1) make it especially suitable for analysis of functional biological preparations. Using computer simulations, we show that the model proposed by Kohler et al. (3) adequately accounts for the main features of the reaction kinetics obtained with this system. PEROXIDASE
ENZYMATIC
CYCLE
To evaluate the peroxidase reaction cycle, the following basic model using two one-electron reduction steps is analyzed (Fig. 1) and is essentially that given in (3). In this scheme two loops are in evidence. Loop 1 is the principal peroxidase catalytic pathway and includes the native enzyme (CO), Compound I (CI), and Compound II (CII). Loop 2 results from the reaction of CII with additional HZOz to form Compound III (CIII) and at least temporarily removes enzyme from the peroxidative cycle. CR1 may then give up oxygen to produce the ferrous enzyme (CIV), which can react with additional HzOz to reform CII. In this way, reversible substrate-dependent inhibition is obtained. In addition, CIII may undergo decomposition to an inactive form which removes enzyme from the peroxidase cycle entirely (3). The complexity of this system rules out an analytical solution for the reaction kinetics so that either steadystate solutions (11) or numerical methods must be used. Although the complete system can be solved using standard numerical methods, the computer time required using a microcomputer can be substantial. This is largely the result of the fact that some rate constants differ by many orders of magnitude, requiring that integrations be carried out with very fine steps (dt). Therefore, as a prac-
AND
MEHLHORN
(feyT-02
c”’ I
Irreversible Forms FIG. 1. Basic model of peroxidase reaction cycle. Two loops have Compound II as the common intermediate. Loop 1 represents the peroxidative path whereas Loop 2 is catalatic, converting HZ02 to H,O and 0,.
tical matter the following simplifying assumptions were made. 1. The reaction rate of the phenoxyl radicals with TOLH is assumed to be instantaneous and irreversible such that for every phenoxyl produced one radical spin label (TOL . )” is obtained. Therefore, the rate of formation of spin probe equals the rate of phenoxyl generation. Since phenol is regenerated with formation of the spin label, and its concentration is typically three orders of magnitude greater than the enzyme concentration, its concentration may be regarded as constant. (We show that these assumptions are justified under Results). 2. It is further assumed that CO, CI and CII are in steady-state. This assumption is reasonable if the reaction rates controlled by kr, k2, and k3 (Fig. 1) are very fast compared to those of the other reactions in the system. Previous results [e.g., (7,9)] indicate that in the presence of electron donors, peroxidative cycling is orders of magnitude faster than conversion rates among the other enzyme intermediates. Our experiments also support this conclusion (see Results). (It should also be noted that the scheme in Fig. 1 can be solved for the full steady-state case; however, this has the undesirable property of effectively eliminating important transients including the initial formation of CIII.) For the peroxidative loop the following results are obtained using the steady-state assumption for CO, CI, and CR: 4 The abbreviation TOL. is used in place of Tempo1 to emphasize the radical form of the spin label and to distinguish it from TOLH, its hydroxylamine derivative.
PEROXIDASE
Co =
ACTIVITY
[C, - (CIII + CIV)]k2k3 phenol klk2H202 + klk3H202 + kzk3 phenol
CI = CII =
[C, - (CIII + CIV)]K1k3H202 klk,H,O,
+ klk3H202 + kzk3 phenol
[C, ~ (CIII + CIV)]k1k2H202 k,k2H202 + klk3Hz02 + kzk3 phenol .
BY ELECTRON
Pal
WI [ICI
Next we have the rate equations for CIII and CIV, and the conservation condition for the total enzyme: dCII1 ~ = k,CIIH202 dt dCIV ~ = k&III dt
- k&III
- k6CIVH20Z
c, = co + CI + CII + CIII + CIV. Finally, the rate expressions for the decomposition HzOz and formation of oxidized spin label are
dHzOz = - [klCO + k&II
____ clt
+ k&IV
dTOL . ~ = k&I phenol + k&II dt
+ k7CI]Hz02
phenol.
WI of
SPIN
[31
Equations [l]-[3] are solved numerically using a Runge-Kutta algorithm to obtain the concentrations of Hz02, spin label (TOL . ), and various forms of the enzyme as a function of time. Parameter fits are obtained using a grid search error minimization protocol comparing the experimental data to simulated solutions (12). METHODS AND MATERIALS ESR. First-derivative ESR spectra were recorded at 22°C using an X-band IBM ER BOOD-SRC spectrometer. Microwave power and modulation amplitude were 10 mW and 1.6-2.0 G, respectively. For kinetic studies, the low field peak of the nitroxide triplet was continuously monitored and used to calculate the nitroxide concentration by comparison with a standard. The ESR stopped-flow apparatus consisted of a computer-controlled solenoid-operated vacuum system as described by Moronne et al. (12). In brief, solutions are combined using a T-mixer connected to the spectrometer sample capillary. Signal recording and flow control are operated using an IBM-AT computer and a DATA Translation 2800 data acquisition board. Acquisition and analysis software were developed using ASYST (MacMillan Publishing), a FORTH-based compiler, and application package. L9pm trap synthesis and standardization. TOLH (1,4-dihydroxy2,2,6,6-tetramethylpiperidine) was synthesized from Tempo1 (2,2,6,6tetramethyl-1-piperidinoxy-4-01) as previously described (13), and stored at ~20°C in concentrations from 150 to 250 mM. Each TOLH stock was assayed by oxidizing a diluted sample containing approximately 100 pM TOLH with 5 mM K,Fe(CN), and quantitated by comparing the line heights with a Tempo1 standard in the same concentration of K:rFe(CN),.
49
All assays were performed Assay conditions and bufferpreparation. in phosphate-buffered saline (PBS) at pH 7.2 and 22°C. HzOz solutions were prepared immediately before use from a 30% stock solution. Tempo1 was purchased from Aldrich. All other reagents were purchased from Sigma. Oxidation of TOLH is catalyzed in solution by transition metal contaminants such as copper and iron. As a consequence, extra precautions were taken to ensure their removal from our solutions. Buffer salts (NaHZP04 and Na2HP0J were dissolved in commercially distilled water, incubated with insoluble phosphateeglass at pH 7.4 until the phosphateglass settled (about 1 week), and then carefully removed from the phosphateeglass. The buffer was then passed through a Chelex 100 (BioRad, Richmond, CA) column which had been washed several times with buffer. Buffers were stored in polypropylene containers to avoid the possibility of absorbing transition metals from glass. These procedures were arrived at empirically using TOLH oxidation assays to monitor the removal of contaminating transition metal ions. In this way baseline oxidation rates of 2.5 mM TOLH in air-equilibrated buffer could be reduced to 0.1% per hour at 22°C and pH 7.4. Oxidation could be completely blocked by inclusion of 20 pM diethylene triaminepentaacetic acid (DTPA) or deferoxamine, compounds that shift the oxidation potential of iron.
RESULTS TOLH Oxidation: Radicals
[2]
RESONANCE
Rapid Scavenging of Phenoxyl
The oxidation of TOLH to its radical form TOL. is slow in the presence of peroxidase and H202 but rapid when the reaction mixture also includes phenol. This is illustrated in Fig. 2, which compares the rate of TOLH oxidation with and without 500 PM phenol measured using ESR stopped-flow. Without phenol, TOLH is very slowly oxidized, indicating that it is not a good substrate for the oxidized peroxidase enzyme intermediates. In contrast, the addition of phenol produces a two order of magnitude stimulation of the oxidation rate. The use of TOL . production as a measure of peroxidase activity requires that reaction of the phenoxyl radical with TOLH be fast compared to other processes such as phenoxyl polymerization. This is illustrated in Fig. 3, which
I 2
60-
i e
40zo-
FIG. 2. Oxidation of TOLH to TOL* with and without phenol. The oxidation of TOLH to its radical form TOL. was measured by ESR stopped-flow. The reaction mixture included 100 pM H202, 1 U HRP/ ml, 2.5 mM TOLH, and the indicated concentrations of phenol.
50
MOORE,
MORONNE,
AND
MEHLHORN
1.2-
z 8 d OBII x, Ii . 0.40
No TOLH
2.5 mM TOLH
i
0
I 50
0
I 100
o.oI 0
I
10
I
20
Time (sec.) FIG. 3. Phenol polymerization with and without TOLH. This figure shows the absorbance change produced at X = 400 nm by peroxidasemediated oxidation of phenol. In the presence of 2.5 mM TOLH, polymer formation is strongly inhibited. The reaction mixture also contained 200 PM phenol, 100 jtM H,O,, and 2 U/ml of HRP.
I 200
Time (sec.)
I
30
1 150
FIG. 4. Ascorbate competition with TOLH for phenoxyl radicals. The reaction mixture included 45 PM ascorbate, 1 U/ml HRP, 100 FM phenol and Hz02, and 2.5 mM TOLH. The data yield an estimate of 1.2 X lo7 Mm’ s-i for the phenoxyl reaction rate with TOLH.
radical form of the spin label gives the following predicted stoichiometry: peroxidase
compares the formation of phenol polymerization products in the presence or absence of 2.5 mM TOLH. The reaction mixtures contained 200 PM phenol, 100 FM HzOz, and 2 U/ml of HRP. The upper curve without the spin trap shows the characteristic absorbance change at 400 nm that results from polymerization of phenol (14). Under these conditions rlj2 is approximately 5 s with a final absorbance change of 1.2 (OD). In the presence of 2.5 mM TOLH (lower curve) there is no increase in absorption, showing that this concentration of TOLH completely suppresses polymer formation. Thus, it appears that the reaction rate of phenoxyl radicals with TOLH is sufficiently rapid to accurately follow radical formation by the peroxidase intermediates. To further confirm the rapid rate of phenoxyl radical reaction with TOLH, an additional control was performed using competition between ascorbate and TOLH. The rate constant reported for reaction of phenoxyl with ascorbate is 6.9 X 10’ M-l 5-l (15). Using a reaction mixture consisting of 1 U/ml HRP, 100 PM phenol and H20z, and 2.5 mM TOLH, it was determined that the rate of TOLH oxidation was 50% inhibited with 45 PM ascorbate (Fig. 4). This leads to an estimate of 1.2 X lo7 M-l spl for the reaction rate of phenoxyl with TOLH. Computer simulations, discussed later, indicate that this rate in conjunction with the 2.5 mM TOLH used throughout these experiments is sufficient to accurately follow the rate of phenoxyl generation and consequently the peroxidase reaction cycle.
HzOz + 2TOLH
From consideration of the reaction cycle shown in Fig. 1, the conversion of Hz02 oxidizing equivalents to the
2TOL.
+ 2H20.
[41
In the presence of sufficient phenol, the oxidation of TOLH by the peroxidase/Hz02 system is rapid and gives close to the theoretical conversion of 2 mol of TOL . per mole of H202. This is shown in Fig. 5, which presents TOLH oxidation curves for various initial H202 concentrations ranging from 0 to 200 PM. The yields are close to the theoretical values, except for the 200 PM case which gave 85% of the expected TOL * . As is discussed in more detail, the loss of conversion efficiency for higher Hz02 concentrations appears to be the result of increased catalatic activity. A slight increase in oxidation rate is seen for increasing initial concentrations of HaOz. This suggests that in the presence of 500 pM phenol, the reaction of phenol with CI and CII approaches the net rate of reaction of H202 with native enzyme (k, - 1 X 107).
0 0
Peroxidase Cycling with Phenol and TOLH
-
I 5
I 10
I
1 20
I 25
I 30
Time’ Tsec.) FIG. 5.
TOLH
oxidation
vs H202. Each trace included 2 U/ml HRP, Buffer included 10 mM phosphate
500 PM phenol, and 2.5 mM TOLH. and 150 mM NaCl at pH 7.2.
PEROXIDASE
Rate of TOL - Production Concentration
Is Proportional
ACTIVITY
BY ELECTRON
SPIN
RESONANCE
to Enzyme
The rate of TOL. formation in the presence of 40 PM H,Oa, 500 PM phenol, and 2.5 mM TOLH is proportional to the HRP concentration. This is shown in Fig. 6, which plots the initial oxidation rate for enzyme concentrations ranging from 0.83 to 830 nM (0.01 to 10 U/ml). We estimate that the minimum enzyme concentrations that can be reliably determined using this approach is about -3 X lo-’ U/ml of HRP (assumes a 50% increase over the background TOLH oxidation rate). The limiting factor largely depends on the rate of TOLH oxidation by HzOz in the presence of contaminating transition metal ions which promote OH. generation. If substrate concentrations (HzOz or phenol) must be minimized to avoid enzyme inactivation or reaction with other system components, a glucose oxidase-HzOz generating system with relatively low phenol concentrations can be used. Figure 7 illustrates the assay of HRP activity with HZO,! provided by 10 mM glucose, 0.2 pg/ml glucose oxidase, atmospheric oxygen, and 40 PM phenol. Again a linear response is obtained for enzyme concentrations in the range of 0.005-0.05 U/ml (0.4-4.0 nM) HRP. At the higher enzyme concentrations, the rate of H202 generation becomes rate limiting. For biological preparations such as cell cultures, this approach ensures low exposure of cells to HzOz and a reduced requirement for phenol or other electron donors to produce stoichiometric conversion of HZOZ to TOL . . In results described elsewhere (20), thyroid peroxidase (TPO) activity in detergent extracts from a Chinese hamster ovarian cell line (CHO-hTP0) transfected with a human TPO gene were evaluated in the range of 0.001-0.005 U/ml using the glucose oxidase system.
HRP (U /ml) FIG. 6. TOLH oxidation vs HRP by ESR stopped-flow. Initial oxidation rates for HRP concentrations from 0.01 to 10 U/ml (0.83-830 nM). Each experiment included 40 PM HzOz, 500 PM phenol, and 2.5 mM TOLH. The basal oxidation rate in the absence of enzyme was subtracted from each value and was one-third the rate of the lowest enzyme concentration.
HRP (U/ml) FIG. ‘7. TOLH oxidation vs HRP using glucose oxidaseeHzOz generator. Initial oxidation rates with HZ02 produced by 10 mM glucose, 0.2 pg/ml of glucose oxidase, and atmospheric oxygen. The oxidation rate eventually saturates as the peroxidase cycling outpaces the rate of Hz02 production.
Enzyme Inhibition
and Its Effects on Stoichiometry
As noted in the introduction, high concentrations of HzOz inhibit peroxidase activity by promoting CIII formation and decomposition of Hz02 (3). Figure 8a shows TOL. accumulation over time with 20, 50, 100, and 200 yM HzOz in the presence of 40 PM phenol and 2 U/ml HRP. Compared to the curves shown in Fig. 5, where 500 PM phenol was used, the TOLH oxidation rates are decreased by about an order of magnitude. In addition, the oxidation rate is increasingly inhibited as the initial concentration of HzOa is increased compared to the slight rise in the initial rates apparent in Fig. 5. Further, these traces show a complex biphasic time dependence which is particularly noticeable for the 100 and 200 PM curves.5 In the first several seconds, there is a rapid rate of oxidation which then slows. This is followed by a second phase during which the oxidation rate accelerates, reaching a maximum shortly before exhaustion of the HzOa. A reversible inactivation process which is consistent with an early accumulation of enzyme in the nonperoxidative path is suggested by these results (Loop 2) while the Hz02 concentration is comparatively high. This behavior can be interpreted as substrate-dependent inhibition producing a dead-end complex (under Results, see Computer Simulations). In further contrast to the data in Fig. 5, significant catalatic activity is clearly manifested for the higher concentrations of H202. The stoichiometry is depressed to only 51% of the theoretical for the 200 PM addition compared to 100% conversion for the 20 PM case. Figure 8b plots the final concentration of TOL. produced versus the initial H,02 concentration. The general trend shows 5 This may be seen more clearly by inclining and sighting along each trace.
the graph at an angle
52
MOORE,
MORONNE,
a.
AND
MEHLHORN
b.
20 ,A4 H,O,
Time (sec.) FIG. 8. TOLH oxidation vs HRP for low phenol. (a) The conditions for this experiment are as described in the legend to Fig. 5 except that 40 pM phenol is used instead. (b) Plot of the final TOL * vs the initial H,O, concentration. The dotted lines in (a) are theoretical fits using computer simulations of the model shown in Fig. 1 with allowance made for the catalatic effect of TOL . (see Computer Simulations under Results). The rate constant for the simulated curves are kl = 1.2 X 10’ Mm’ s-l, k2 = 2 X 10s Mm' SC', k3 = 1.2 X lo5 Me' s-l, k4 = 650 Mm' s-l, k, = 0.04 s-l, ks = 5 X lo4 M-' s-l, k7 = 500 M-' s-l, $ = 20 M-l s-l HRP-‘.
a relative loss of oxidizing equivalents for increased initial concentrations of HzOz. This result is in keeping with the model proposed by Kohler et al. (3), who found that the reconversion of CIII to native enzyme for LPO and TPO generated approximately 1 mol of O2 for every 2 mol of HaOz consumed. Presumably, this results from the release of O2 by CIII to give the ferrous enzyme (CIV), which then combines with H202 to give CR. However, as is discussed (under Computer Simulations), it appears that this mechanism is unable to account for the large amount of H202 decomposition observed under conditions of low phenol concentrations. Recent results (16) indicate that TOL * is able to promote catalatic activity in the presence of heme proteins and is able to account for the discrepancy. Effects of Phenol on Substrate-Dependent
Inhibition
The HzOz-dependent inhibition of the peroxidase enzyme activity is a function of the electron donor concentration [e.g., Fig. 5 vs Figs. 8a and 8b, this paper; and Ref. (7)]. Figure 9 shows the effects of increasing phenol concentration on the initial TOLH oxidation rates mediated by HRP and LPO. The rate of oxidation begins to saturate for both enzymes as the phenol concentration is raised. Phenol produces much the same increase in the LPOmediated rate of TOLH oxidation as it does for HRP. However, for the same unit concentration of enzyme (pyrogallol units) and the same phenol concentration, LPO appears to have about twice the activity of HRP. Interestingly, in contrast to the HRP results, biphasic kinetics were not observed with LPO even at low phenol concentrations (40 PM). This suggests that the rate of CIII formation for LPO is either slower than that for HRP or that the phenol enhancement of the peroxidase cycling is faster, producing less CIII and CIV.
Computer Simulations In order to validate the reaction cycle in Fig. 1, two notable features of the experimental data must be explained by the proposed model. First, the model must be able to generate the biphasic characteristics of the TOLH oxidation curves evident in samples containing comparatively low concentrations of phenol and large concentrations of HzOZ (Fig. 8). Second, the model must account for the decreasing conversion efficiency of HzOz equivalents to TOL * under the same conditions. Computer simulations using numerical solutions to Eqs. [l-3] indicate that biphasic kinetics of TOLH oxidation can be produced by the Kohler model. Figures 10a and lob show the predicted time-dependent concentrations of TOL a, HtOa, and enzyme components in a system with 40 PM phenol, 200 PM H202, and 2 U/ml HRP (same as in Fig. 8). Two cases are considered: (i) the basic Kohler model as depicted in Fig. 1 (solid lines) and (ii) a modified
--c) Q-----------------30
-I
Kl
.’
*’
.-
.-
.’
.*’
, ,’ P’ I’ I’ .’ t A.
,&----
_--- --
. o- LPO A- HRP
--*-------------------
FIG. 9. Initial TOLH oxidation rates vs phenol. Phenol stimulation of TOLH oxidation for HRP and LPO. Enzyme = 1 U/ml, 50 pM H,Oa, 2.5 mM TOLH, pH 7.2, at 22°C.
PEROXIDASE
ACTIVITY
BY ELECTRON
SPIN
53
RESONANCE
a.
200 f
HA 150
d.
b.
260
360
Time (sec.) FIG. 10. (a-d) Simulations based on the model in Fig. 1. (a) Predicted concentrations of HZ02 and TOL. in the case of 40 /.tM phenol (solid and dashed lines represent unmodified and modified models respectively); (b) corresponding enzyme concentrations. (c and d) The same quantities, but calculated for the case of 500 PM phenol. In each case the following rate constants were used: k, = 1.2 X lo7 Mm1 s-l, k2 = 2 X lo6 Me’ se’, k, = 1.1 X lo5 M ’ s I, k, = 250 M-’ s ‘, k, = 0.05 s-l, k, = 5 X lo4 M-’ s-i, k, = 500 M-’ s-i, k8 = 20 M-’ s-i HRP.
version which includes correction for TOL * -dependent catalatic activity (dashed lines). Both models demonstrate the characteristic kinetic behavior associated with substrate inhibition. The initial rates of TOLH oxidation are comparatively rapid but then decline. Thereafter, the oxidation rates accelerate, reaching a maximum shortly before exhaustion of the H202. As shown in Fig. lob, this behavior is reflected in the concentration of nonperoxidative enzyme (CIII + CIV) that at first rapidly increases coincident with the early inactivation of peroxidase activity. Subsequently, as HzOz is consumed, the concentrations of CIII and CIV decrease, gradually restoring activity. Although the biphasic kinetics are well represented, the unmodified model predicts a yield very close to the maximum value of 2 TOL . per H,O, even for 40 PM phenol. This is not consistent with the experimental results in Fig. 8 that indicate substantial catalatic action with loss of H,O,. According to the simulation only 5 j.rM of 200 PM H,O, is lost to catalatic activity (Loop 2); this is in contrast to experimental results showing 95 PM H,Oz destroyed (Fig. 8). The computer result is obtained despite the fact that the value of k4 used in the simulation was 10 times higher than previous literature values (4, 17). Even with the higher rate constant, cycling through Loop
2 is apparently too slow to account for the observed destruction of H,Oz. Recent experiments with heme proteins have shown that nitroxides can promote catalatic action (16). Nitroxides are not consumed during this process suggesting the following: peroxidase
2H202 + TOL * -
ks
2H,0+02+TOL..
(5)
Without attempting to specify the details of this process, oximetry data yield an estimate of 20 M-’ s-l per unit HRP for k8 under the experimental conditions used in the current studies. Including this reaction and the estimated value for k8 in the computer model produces the dotted line traces shown in Figs. 10a and lob. In this case, the loss of Hz02 is very close to the experimental result. The adequacy of the model is further illustrated by the dotted theoretical lines shown in Fig. Ba. Each of the theoretical curves was calculated based on a fit to the experimental data for the 200 PM H,O, trace. The data for each concentration of Hz02 agree reasonably well with the calculated curves, indicating self-consistency. In addition, the disappearance of biphasic behavior for the
54
MOORE,
MORONNE,
lowest concentrations of HzOz, as well as the near stoichiometric conversion of HaOz to TOL . , is also properly indicated. Figures 1Oc and 10d show the predicted rate of TOLH oxidation and the various enzyme forms when the electron donor concentration (phenol) is increased to 500 pM (as in Fig. 5). In this case, the TOLH oxidation rate is increased an order of magnitude (note the time scale), biphasic kinetics are no longer apparent, and the catalatic decomposition of Hz02 is greatly reduced. Again these data are consistent with the experimental results in Figs. 5 and 8. DISCUSSION
Measurement
of H202 and Peroxidase Activity
by ESR
We have shown that ESR methods can be used to measure trace activities of peroxidase enzymes (e.g., HRP activities as low as 0.003 U/ml pyrrogallol units). The assay is suitable for peroxidase enzymes that can be obtained free of reducing substrates. If this condition is fulfilled, then a substrate concentration-dependent stimulation of TOLH oxidation in the presence of hydrogen peroxide is sufficient to demonstrate the presence of peroxidase activity. This presumes that the oxidized substrates are capable of converting TOLH to its radical form [reduction potential = 0.3 V for the nearly identical TEMPOH, Ref. (IS)]. In this regard, the peroxidase oxidation products of biologically important substrates such as phenols and halides are potent TOLH oxidants. Further, the rates of reaction with TOLH are sufficiently fast that production of TOL . is an accurate measure of the rate of production of the oxidized intermediate. The fact that the peroxidase substrate is regenerated upon reaction with TOLH means that its concentration is effectively clamped, helping to maintain a steady rate of TOLH oxidation and enzyme cycling. This is a particular analytical advantage since the evaluation of activity is not limited to precise extrapolations of initial rates which are often complicated by the presence of noise and other artifacts such as mixing transients. Another advantage is that the ESR approach is sparing of biological materials. For example, with a typical 20-4 ESR sample the activity of as little as 6 pmol of enzyme can be measured. This sensitivity can be increased by an order of magnitude using a loop gap resonator which requires a sample size less than 2 ~1 (19). The efficacy of the ESR assay suggests its utility in systems where sensitivity and response time are important, such as samples with either very low or high activity. In the case of the latter, reaction rates on the millisecond time scale can be measured with ESR stopped-flow. For samples with low activity the accumulation of submicromolar concentrations of the stable radical product TOL . can be readily followed free of optical interference. HzOz and organic hydroperoxides capable of forming CI
AND
MEHLHORN
can be also assayed with peroxidase and an excess of an electron donor such as phenol so that peroxide is stoichiometrically converted to TOL * (Fig. 5). In addition, HZOz produced enzymatically can be quantitated using high peroxidase and electron donor concentrations. In this instance the rate of TOL . production will be determined by the rate of HzOz generation (e.g., Fig. 7). The basic constraint of the ESR assay and peroxidase assays in general is that conditions be chosen to minimize enzyme inactivation by conversion either to inactive but reversible forms such as CIII and CIV or to fully degraded enzyme. Inactivation is promoted by prolonged exposure to high concentrations of H202. As a consequence, peroxidase activity is best measured with low levels of Hz02 ( 0.5 mM produce simple kinetics and give nearly theoretical conversion of HzOz to TOL . . There have been a number of recent studies on the formation of CIII by high levels of HzOz and the reconversion of CIII to native enzyme (3, 4, 7, 9, 14, 17). In regard to the ESR assay, pertinent mechanisms identified for Compound III formation include (i) the direct reaction of HzOZ with CII (3, 4) and (ii) addition of superoxide anion to ferriperoxidase (4, 22, 23). In the ESR system, the former reaction seems highly probable and is consistent with increasing inhibition of TOLH oxidation with increasing HZOz. However, the direct participation of superoxide seems unlikely. It has been determined that reduced nitroxides similar to TOLH scavenge superoxide, producing TOL . and HzOz. Rosen et al. (24) determined a rate constant of -lo3 M-l s--l for superoxide and TEMPOH, a close analog of TOLH. Thus, the relatively high concentration of TOLH (2.5 mM) used in these experiments argues against the participation of freely diffusible superoxide. In addition, nitroxides such as TOL . rapidly form superoxide adducts that are potent oxidants of thiols and probably TOLH. The rate constant for adduct formation is estimated to be 3 X 10” M-’ s-’ (25). As a consequence, TOL * produced during TOLH oxidation is most likely to increase superoxide scavenging. Although it appears unlikely that superoxide contributes to CIII formation in the ESR-TOLH system, analyses of experiments using low phenol (40 PM, Fig. 8) indicate that the rate of CIII formation is enhanced compared to spectrophotometric measurements done in the absence of electron donors (4, 17). To fit data such as those in Fig. 8, it. was found that k, (Fig. 1) needed to be 5-20 times higher than the previous values of 20-25 M ’ so) (4, 17). Simulations using the slower literature value did not produce a sufficient rate of peroxidase inactivation to account for either the initial inactivation phase or the subsequent recovery of activity; in other words, too little enzyme is directed through Loop 2. This result suggests that phenol, TOLH, or TOL. is able to directly or indirectly facilitate the conversion of CII to CIII. In the case of phenol, Ma and Rokita (14) determined that the rate of irreversible peroxidase inactivation of HRP by Hz02 was increased an order of magnitude when phenol was included in the reaction mixture. Since it has been argued that CIII is an intermediate that leads to irreversible inactivation (3), this may indicate that phenol or its radical products facilitate formation of CIII. The
SPIN
55
RESONANCE
involvement of phenoxyl radicals in the ESR system seems unlikely since the steady-state concentration is expected to be very low given the rapid scavenging provided by TOLH. The possibility that TOLH is able to promote CIII formation also seems remote. Experiments comparing 0.1, 0.5, and 2.5 mM TOLH yielded the same overall rates of TOLH oxidation, suggesting that formation of CIII is unaffected by TOLH. As discussed earlier, TOL * promotes the catalatic activity of heme proteins including HRP. The exact mechanism remains to be elucidated. However, one possible scenario involves the oxidation of TOL . to the immonium oxene cation [TOL+, Ref. (26)] by CI, CII, or phenoxyl. The oxene compound may oxidize H202 to give water and oxygen, thus acting as a catalase, or might give an adduct capable of converting native enzyme to CIII. When the initial TOL . concentration was increased from 90 to 450 PM in a mixture consisting of 1 U/ml HRP, 100 PM HzOz, and 0.5 mM phenol, a 20% drop in conversion efficiency and peroxidase activity (oxidation rate) resulted. The fact that the activity declined indicates that high TOL . may promote CIII formation. ACKNOWLEDGMENTS We thank Dr. Judith Klinman of the Department of Chemistry, UC Berkeley, for helpful discussions. This work was supported by NIH Grant AG 04818, the Cigarette and Tobacco Surtax Fund of the State of California through the Tobacco-Related Disease Research Program of the University of California, and Office of Energy Research, through the Department of Energy under Contract DE-AC03-76SF00098.
REFERENCES 1. Dunford, H. B. (1991) in Peroxidases in Chemistry and Biology (Everse, J., Everse, K. E., and Grisham, M. C., Eds.), Vol 2, CRC Press, Boca Raton, FL. 2. Kohler, H., and Jenzer, H. (1989) Free Rad. Biol. Med. 6, 323-339. 3. Kohler, H., Taurog, A., and Dunford, Biophys. 264(2),438-449. 4. Nakajima, 2581.
R., andyamazaki,
H. 8. (1988) Arch. Biochem.
I. (1987) J. Biol. Chem. 262(6),
2576-
5. Nichols, P. (1966) in Hemes and Hemoproteins (Chance, B., Estabrook, R., and Yonetani, T., Eds.), pp. 307-318, Academic Press, New York. 6. Thomas, E. L., Bozeman, P. M., and Learn, D. B. (1991) in Peroxidases in Chemistry and Biology (Everse, J., et al., Eds.), Vol. I, CRC Press, Boca Raton, FL. I. Amao, M. B., Acosta, M., de1 Rio, J. A., and Garcia-Canovas, F. (1989) Biochim. Biophys. Acta 1038, 85-89. 8. Yamazaki, I., and Piette, L. H. (1963) Biochim. Biophys. Acta 77, 47-64. 9. Jenzer, H., Kohler, H., and Broger, C. (1987) Arch. Biochem. Biophys. 258(2), 381-390. 10. Hosoya, T., Sato, I., Hiyama, Y., Yoshimura, H., Niimi, H., and Tarutani, 0. (1985) J. Biochem. 98, 637-647. 11. Bardsley, W. G. (1985) in The Lactoperoxidase System: Chemistry and Biological Significance (Pruitt, K. M., and Tenovuo, J. O., Eds.), Dekker, New York.
56
MOORE,
MORONNE,
12. Moronne, M. M., Mehlhorn, R. J., Miller, M. P., Ackerson, L. C., and Macey, R. I. (1990) J. Memb. Biol. 115, 31-40. 13. Prolla, T. A., and Mehlhorn, R. J. (1990) Free Rd. Res. Commun. 9,135-146. 14. Ma, X., and Rokita, S. E. (1988) Biochem. Biophys. Res. Commun. 157(l), 170-175. 15. Schuler, R. H. (1977) Radiat. Res. 69, 417-433. 16. Mehlhorn, R., and Swanson, C. E. (1992) Free Rd. Res. Commun., in press. 1’7. Adediran, S. A., and Lambeir, A. (1989) Eur. J. Biochem. 186,571I Fil6. 18. Rosen, G. M., Rauckman,
E. J., and Hanck, K. W. (1977) Toricol.
Lett. 1, 71-74. 19. Hubbell, W. L., Froncisz, W., and Hyde, J. S. (1987) Reu. Sci. Znstrum. 58, 1879-1896.
AND
MEHLHORN
20. Moore, K. L., (1990) PhD dissertation, Berkeley. 21. Jenzer, H., and Kohler,
University
H. (1986) Biochem. Rio&y.
of California, Res. Commun.
139(1),327-332. 22. Wittenberg, J. B., Noble, R. W., Wittenberg, B. A., Antonini, E., Brunori, M., and Wyman, J. (1967) J. Biol. Chem. 242, 626-634. 23. Metodiewa, D., and Dunford, 272(l), 245-253.
H. B. (1989) Arch. Biochem. Biophys.
24. Rosen, G. M., Finkelstein, E., and Rauckman, Biochem. Biophys. 215, 367-378.
E. J. (1982) Arch.
25. Finkelstein, E., Rosen, G. M., and Rauckman, E. J. (1984) Biochim. Biophys. Acta 802,90-98. 26. Rozantzev, E. G. (1970) in Free Nitroxyl Plenum, New York.
Radicals (Ulrich,
H., Ed.),
OF BIOCHEMISTRY
Vol. 299, No. 1, November
AND
BIOPHYSICS
15, pp. 47-56,
1992
Electron Spin Resonance Study of Peroxidase Activity and Kinetics’ Katherine
L. Moore,
Division
of Energy
Received
April
Mario
M. Moronne,
and Environment,
22, 1992. and in revised
Lawrence
form
July
and Rolf J. Mehlhorn’ Berkeley
Laboratory,
Horseradish peroxidase (HRP)3 and lactoperoxidase (LPO) are typical of peroxidase enzymes whose catalytic ’ The U.S. Government’s right to retain a nonexclusive royalty-free license in and to the copyright covering this paper, for governmental purposes, is acknowledged. ’ To whom correspondence should be addressed at Division of Energy and Environment, M/S 7O-193A, Lawrence Berkeley Laboratory, 1 Cyclotron Road, Berkeley, CA 94720. 3 Abbreviations used: HRP, horseradish peroxidase; LPO, lactoperoxidase; ABTS, 2,2’-azinobis(3-ethylbenzthiazoline-6-sulfonic acid; TOLH, 1,4-dihydroxy-2,2,6,6-tetramethylpiperadine; Tempol, 2,2,6,6tetramethyl-l-piperidinoxy-4-01; PBS, phosphate-buffered saline; DTPA, diethylene triaminepentaacetic acid; TPO, thyroid peroxidase.
$5.00
Copyright 8 1992 by Academic Press, Inc. All rights of reproduction in any form reserved.
California
94720
22, 1992
An electron spin resonance (ESR) assay has been developed for peroxidase activity. The assay measures the formation of the paramagnetic nitroxide Tempo1 from the oxidation of its hydroxylamine derivative (TOLH) by short-lived radicals produced by peroxidase cycle intermediates, Compounds I and II. Using phenol as a peroxidase electron donor, the ESR approach is suitable for measurements of peroxidase activity (aO.003 U/ml) and micromolar quantities of HzOz in sample sizes as small as 2 ~1. In addition, the ESR method can be used to continuously monitor activity in cell suspensions and other media that are susceptible to optical artifacts. The high membrane permeability of TOLH also makes it possible to estimate peroxidase activity in membrane-enclosed compartments, provided that TOLH oxidation rates can be stimulated with exogenous peroxidase reductants, e.g., phenol. Analysis of TOLH oxidation rates under conditions of low electron donor concentrations and high concentrations of HzOz also shows clear indications of substrate-dependent inhibition and increased catalatic activity. Computer simulations indicate that the results obtained are consistent with the peroxidase reaction scheme proposed by Kohler et al. (1988, Arch. Biochem. Biophys. 264, 438-449) modified to correct for a nitroxide dependent stimulation of peroxidase catalytic activity. ‘tt 1992 Academic Press, Inc.
oow9861/92
Berkeley,
cycle involves a number of distinct intermediates with differing oxidation states of the active iron center (for reviews see (1) and (2)). Native enzyme (ferriperoxidaseFe3+) reacts with HzOz to form Compound I (Fe5+), which may then return to the native enzyme by either a single two-electron reduction or by two one-electron steps. The source of donor electrons determines the path and rate of Compound I conversion back to native enzyme. Compound I is rapidly reduced by iodide in a single two-electron process (3). At a much slower rate, H,Oz may also serve as a two-electron donor largely accounting for the modest catalatic activity of peroxidases (3, 4). In the absence of an exogenous electron donor, Compound I can return to native enzyme in two relatively slow oneelectron reactions with reducing groups contained within the enzyme. For HRP, this process can proceed through 10 or more cycles (5). Exogenous one-electron donors (e.g., phenol) greatly increase enzyme turnover by facilitating transformation of Compound I to Compound II and Compound II back to native enzyme. In fact, a variety of inorganic and organic electron donors, including sulfur compounds and halogenated and nonhalogenated phenols, are suitable substrates for these peroxidase intermediates (2, 6). Various studies (3, 4, 7) have shown that H,OZ, in addition to serving as a substrate, can inhibit the catalytic cycle by reacting with Compound II to form Compound III, which is outside the peroxidatic loop (see Fig. 1). This has been proposed to occur by two different pathways, one of which depends on the formation of superoxide (4, 8). Subsequent to formation, Compound III can release oxygen to yield the ferrous enzyme, which further reacts with Hz02 to regenerate Compound II (3,9). Alternatively, Compound III may irreversibly decompose, resulting in a permanent loss of peroxidase activity (3). In the presence of appropriate electron donors such as 2,2’-azinobis(3ethylbenzthiazoline-6-sulfonic acid) (ABTS), the formation of Compound III can be reduced (7), thereby decreasing enzyme inactivation when exposed to high con47
48
MOORE,
MORONNE,
centrations of HzOz. This last feature is important in the optimal design of systems to monitor peroxidase activity in that conditions that conserve the enzyme can be chosen. In this paper, we present the results of an electron spin resonance (ESR) study of the peroxidase reaction cycle. In particular, the reaction cycle is evaluated using an ESR assay for the one-electron oxidation products of the peroxidase . HzOz complex using the reduced hydroxylamine derivative of the spin probe Tempo1 (TOLH). The approach taken depends on the capacity of Compounds I and II to rapidly oxidize phenolic substrates, producing radical intermediates which in turn oxidize the reduced nitroxide to its radical form. In this process, the phenol is regenerated and the accumulation of the nitroxide monitored by ESR spectrometry. With this approach, peroxidase activity > 0.003 U/ml is readily measured. Although accepted techniques for evaluation of peroxidase activity already exist (e.g., the guaiacol assay, Ref. (lo)), previous methods have largely depended on optical measurements that are unsuitable for many biological preparations such as turbid cell or microsomal suspensions. ESR is essentially free of optical artifacts, and its high sensitivity and small sample requirements (2-20 ~1) make it especially suitable for analysis of functional biological preparations. Using computer simulations, we show that the model proposed by Kohler et al. (3) adequately accounts for the main features of the reaction kinetics obtained with this system. PEROXIDASE
ENZYMATIC
CYCLE
To evaluate the peroxidase reaction cycle, the following basic model using two one-electron reduction steps is analyzed (Fig. 1) and is essentially that given in (3). In this scheme two loops are in evidence. Loop 1 is the principal peroxidase catalytic pathway and includes the native enzyme (CO), Compound I (CI), and Compound II (CII). Loop 2 results from the reaction of CII with additional HZOz to form Compound III (CIII) and at least temporarily removes enzyme from the peroxidative cycle. CR1 may then give up oxygen to produce the ferrous enzyme (CIV), which can react with additional HzOz to reform CII. In this way, reversible substrate-dependent inhibition is obtained. In addition, CIII may undergo decomposition to an inactive form which removes enzyme from the peroxidase cycle entirely (3). The complexity of this system rules out an analytical solution for the reaction kinetics so that either steadystate solutions (11) or numerical methods must be used. Although the complete system can be solved using standard numerical methods, the computer time required using a microcomputer can be substantial. This is largely the result of the fact that some rate constants differ by many orders of magnitude, requiring that integrations be carried out with very fine steps (dt). Therefore, as a prac-
AND
MEHLHORN
(feyT-02
c”’ I
Irreversible Forms FIG. 1. Basic model of peroxidase reaction cycle. Two loops have Compound II as the common intermediate. Loop 1 represents the peroxidative path whereas Loop 2 is catalatic, converting HZ02 to H,O and 0,.
tical matter the following simplifying assumptions were made. 1. The reaction rate of the phenoxyl radicals with TOLH is assumed to be instantaneous and irreversible such that for every phenoxyl produced one radical spin label (TOL . )” is obtained. Therefore, the rate of formation of spin probe equals the rate of phenoxyl generation. Since phenol is regenerated with formation of the spin label, and its concentration is typically three orders of magnitude greater than the enzyme concentration, its concentration may be regarded as constant. (We show that these assumptions are justified under Results). 2. It is further assumed that CO, CI and CII are in steady-state. This assumption is reasonable if the reaction rates controlled by kr, k2, and k3 (Fig. 1) are very fast compared to those of the other reactions in the system. Previous results [e.g., (7,9)] indicate that in the presence of electron donors, peroxidative cycling is orders of magnitude faster than conversion rates among the other enzyme intermediates. Our experiments also support this conclusion (see Results). (It should also be noted that the scheme in Fig. 1 can be solved for the full steady-state case; however, this has the undesirable property of effectively eliminating important transients including the initial formation of CIII.) For the peroxidative loop the following results are obtained using the steady-state assumption for CO, CI, and CR: 4 The abbreviation TOL. is used in place of Tempo1 to emphasize the radical form of the spin label and to distinguish it from TOLH, its hydroxylamine derivative.
PEROXIDASE
Co =
ACTIVITY
[C, - (CIII + CIV)]k2k3 phenol klk2H202 + klk3H202 + kzk3 phenol
CI = CII =
[C, - (CIII + CIV)]K1k3H202 klk,H,O,
+ klk3H202 + kzk3 phenol
[C, ~ (CIII + CIV)]k1k2H202 k,k2H202 + klk3Hz02 + kzk3 phenol .
BY ELECTRON
Pal
WI [ICI
Next we have the rate equations for CIII and CIV, and the conservation condition for the total enzyme: dCII1 ~ = k,CIIH202 dt dCIV ~ = k&III dt
- k&III
- k6CIVH20Z
c, = co + CI + CII + CIII + CIV. Finally, the rate expressions for the decomposition HzOz and formation of oxidized spin label are
dHzOz = - [klCO + k&II
____ clt
+ k&IV
dTOL . ~ = k&I phenol + k&II dt
+ k7CI]Hz02
phenol.
WI of
SPIN
[31
Equations [l]-[3] are solved numerically using a Runge-Kutta algorithm to obtain the concentrations of Hz02, spin label (TOL . ), and various forms of the enzyme as a function of time. Parameter fits are obtained using a grid search error minimization protocol comparing the experimental data to simulated solutions (12). METHODS AND MATERIALS ESR. First-derivative ESR spectra were recorded at 22°C using an X-band IBM ER BOOD-SRC spectrometer. Microwave power and modulation amplitude were 10 mW and 1.6-2.0 G, respectively. For kinetic studies, the low field peak of the nitroxide triplet was continuously monitored and used to calculate the nitroxide concentration by comparison with a standard. The ESR stopped-flow apparatus consisted of a computer-controlled solenoid-operated vacuum system as described by Moronne et al. (12). In brief, solutions are combined using a T-mixer connected to the spectrometer sample capillary. Signal recording and flow control are operated using an IBM-AT computer and a DATA Translation 2800 data acquisition board. Acquisition and analysis software were developed using ASYST (MacMillan Publishing), a FORTH-based compiler, and application package. L9pm trap synthesis and standardization. TOLH (1,4-dihydroxy2,2,6,6-tetramethylpiperidine) was synthesized from Tempo1 (2,2,6,6tetramethyl-1-piperidinoxy-4-01) as previously described (13), and stored at ~20°C in concentrations from 150 to 250 mM. Each TOLH stock was assayed by oxidizing a diluted sample containing approximately 100 pM TOLH with 5 mM K,Fe(CN), and quantitated by comparing the line heights with a Tempo1 standard in the same concentration of K:rFe(CN),.
49
All assays were performed Assay conditions and bufferpreparation. in phosphate-buffered saline (PBS) at pH 7.2 and 22°C. HzOz solutions were prepared immediately before use from a 30% stock solution. Tempo1 was purchased from Aldrich. All other reagents were purchased from Sigma. Oxidation of TOLH is catalyzed in solution by transition metal contaminants such as copper and iron. As a consequence, extra precautions were taken to ensure their removal from our solutions. Buffer salts (NaHZP04 and Na2HP0J were dissolved in commercially distilled water, incubated with insoluble phosphateeglass at pH 7.4 until the phosphateglass settled (about 1 week), and then carefully removed from the phosphateeglass. The buffer was then passed through a Chelex 100 (BioRad, Richmond, CA) column which had been washed several times with buffer. Buffers were stored in polypropylene containers to avoid the possibility of absorbing transition metals from glass. These procedures were arrived at empirically using TOLH oxidation assays to monitor the removal of contaminating transition metal ions. In this way baseline oxidation rates of 2.5 mM TOLH in air-equilibrated buffer could be reduced to 0.1% per hour at 22°C and pH 7.4. Oxidation could be completely blocked by inclusion of 20 pM diethylene triaminepentaacetic acid (DTPA) or deferoxamine, compounds that shift the oxidation potential of iron.
RESULTS TOLH Oxidation: Radicals
[2]
RESONANCE
Rapid Scavenging of Phenoxyl
The oxidation of TOLH to its radical form TOL. is slow in the presence of peroxidase and H202 but rapid when the reaction mixture also includes phenol. This is illustrated in Fig. 2, which compares the rate of TOLH oxidation with and without 500 PM phenol measured using ESR stopped-flow. Without phenol, TOLH is very slowly oxidized, indicating that it is not a good substrate for the oxidized peroxidase enzyme intermediates. In contrast, the addition of phenol produces a two order of magnitude stimulation of the oxidation rate. The use of TOL . production as a measure of peroxidase activity requires that reaction of the phenoxyl radical with TOLH be fast compared to other processes such as phenoxyl polymerization. This is illustrated in Fig. 3, which
I 2
60-
i e
40zo-
FIG. 2. Oxidation of TOLH to TOL* with and without phenol. The oxidation of TOLH to its radical form TOL. was measured by ESR stopped-flow. The reaction mixture included 100 pM H202, 1 U HRP/ ml, 2.5 mM TOLH, and the indicated concentrations of phenol.
50
MOORE,
MORONNE,
AND
MEHLHORN
1.2-
z 8 d OBII x, Ii . 0.40
No TOLH
2.5 mM TOLH
i
0
I 50
0
I 100
o.oI 0
I
10
I
20
Time (sec.) FIG. 3. Phenol polymerization with and without TOLH. This figure shows the absorbance change produced at X = 400 nm by peroxidasemediated oxidation of phenol. In the presence of 2.5 mM TOLH, polymer formation is strongly inhibited. The reaction mixture also contained 200 PM phenol, 100 jtM H,O,, and 2 U/ml of HRP.
I 200
Time (sec.)
I
30
1 150
FIG. 4. Ascorbate competition with TOLH for phenoxyl radicals. The reaction mixture included 45 PM ascorbate, 1 U/ml HRP, 100 FM phenol and Hz02, and 2.5 mM TOLH. The data yield an estimate of 1.2 X lo7 Mm’ s-i for the phenoxyl reaction rate with TOLH.
radical form of the spin label gives the following predicted stoichiometry: peroxidase
compares the formation of phenol polymerization products in the presence or absence of 2.5 mM TOLH. The reaction mixtures contained 200 PM phenol, 100 FM HzOz, and 2 U/ml of HRP. The upper curve without the spin trap shows the characteristic absorbance change at 400 nm that results from polymerization of phenol (14). Under these conditions rlj2 is approximately 5 s with a final absorbance change of 1.2 (OD). In the presence of 2.5 mM TOLH (lower curve) there is no increase in absorption, showing that this concentration of TOLH completely suppresses polymer formation. Thus, it appears that the reaction rate of phenoxyl radicals with TOLH is sufficiently rapid to accurately follow radical formation by the peroxidase intermediates. To further confirm the rapid rate of phenoxyl radical reaction with TOLH, an additional control was performed using competition between ascorbate and TOLH. The rate constant reported for reaction of phenoxyl with ascorbate is 6.9 X 10’ M-l 5-l (15). Using a reaction mixture consisting of 1 U/ml HRP, 100 PM phenol and H20z, and 2.5 mM TOLH, it was determined that the rate of TOLH oxidation was 50% inhibited with 45 PM ascorbate (Fig. 4). This leads to an estimate of 1.2 X lo7 M-l spl for the reaction rate of phenoxyl with TOLH. Computer simulations, discussed later, indicate that this rate in conjunction with the 2.5 mM TOLH used throughout these experiments is sufficient to accurately follow the rate of phenoxyl generation and consequently the peroxidase reaction cycle.
HzOz + 2TOLH
From consideration of the reaction cycle shown in Fig. 1, the conversion of Hz02 oxidizing equivalents to the
2TOL.
+ 2H20.
[41
In the presence of sufficient phenol, the oxidation of TOLH by the peroxidase/Hz02 system is rapid and gives close to the theoretical conversion of 2 mol of TOL . per mole of H202. This is shown in Fig. 5, which presents TOLH oxidation curves for various initial H202 concentrations ranging from 0 to 200 PM. The yields are close to the theoretical values, except for the 200 PM case which gave 85% of the expected TOL * . As is discussed in more detail, the loss of conversion efficiency for higher Hz02 concentrations appears to be the result of increased catalatic activity. A slight increase in oxidation rate is seen for increasing initial concentrations of HaOz. This suggests that in the presence of 500 pM phenol, the reaction of phenol with CI and CII approaches the net rate of reaction of H202 with native enzyme (k, - 1 X 107).
0 0
Peroxidase Cycling with Phenol and TOLH
-
I 5
I 10
I
1 20
I 25
I 30
Time’ Tsec.) FIG. 5.
TOLH
oxidation
vs H202. Each trace included 2 U/ml HRP, Buffer included 10 mM phosphate
500 PM phenol, and 2.5 mM TOLH. and 150 mM NaCl at pH 7.2.
PEROXIDASE
Rate of TOL - Production Concentration
Is Proportional
ACTIVITY
BY ELECTRON
SPIN
RESONANCE
to Enzyme
The rate of TOL. formation in the presence of 40 PM H,Oa, 500 PM phenol, and 2.5 mM TOLH is proportional to the HRP concentration. This is shown in Fig. 6, which plots the initial oxidation rate for enzyme concentrations ranging from 0.83 to 830 nM (0.01 to 10 U/ml). We estimate that the minimum enzyme concentrations that can be reliably determined using this approach is about -3 X lo-’ U/ml of HRP (assumes a 50% increase over the background TOLH oxidation rate). The limiting factor largely depends on the rate of TOLH oxidation by HzOz in the presence of contaminating transition metal ions which promote OH. generation. If substrate concentrations (HzOz or phenol) must be minimized to avoid enzyme inactivation or reaction with other system components, a glucose oxidase-HzOz generating system with relatively low phenol concentrations can be used. Figure 7 illustrates the assay of HRP activity with HZO,! provided by 10 mM glucose, 0.2 pg/ml glucose oxidase, atmospheric oxygen, and 40 PM phenol. Again a linear response is obtained for enzyme concentrations in the range of 0.005-0.05 U/ml (0.4-4.0 nM) HRP. At the higher enzyme concentrations, the rate of H202 generation becomes rate limiting. For biological preparations such as cell cultures, this approach ensures low exposure of cells to HzOz and a reduced requirement for phenol or other electron donors to produce stoichiometric conversion of HZOZ to TOL . . In results described elsewhere (20), thyroid peroxidase (TPO) activity in detergent extracts from a Chinese hamster ovarian cell line (CHO-hTP0) transfected with a human TPO gene were evaluated in the range of 0.001-0.005 U/ml using the glucose oxidase system.
HRP (U /ml) FIG. 6. TOLH oxidation vs HRP by ESR stopped-flow. Initial oxidation rates for HRP concentrations from 0.01 to 10 U/ml (0.83-830 nM). Each experiment included 40 PM HzOz, 500 PM phenol, and 2.5 mM TOLH. The basal oxidation rate in the absence of enzyme was subtracted from each value and was one-third the rate of the lowest enzyme concentration.
HRP (U/ml) FIG. ‘7. TOLH oxidation vs HRP using glucose oxidaseeHzOz generator. Initial oxidation rates with HZ02 produced by 10 mM glucose, 0.2 pg/ml of glucose oxidase, and atmospheric oxygen. The oxidation rate eventually saturates as the peroxidase cycling outpaces the rate of Hz02 production.
Enzyme Inhibition
and Its Effects on Stoichiometry
As noted in the introduction, high concentrations of HzOz inhibit peroxidase activity by promoting CIII formation and decomposition of Hz02 (3). Figure 8a shows TOL. accumulation over time with 20, 50, 100, and 200 yM HzOz in the presence of 40 PM phenol and 2 U/ml HRP. Compared to the curves shown in Fig. 5, where 500 PM phenol was used, the TOLH oxidation rates are decreased by about an order of magnitude. In addition, the oxidation rate is increasingly inhibited as the initial concentration of HzOa is increased compared to the slight rise in the initial rates apparent in Fig. 5. Further, these traces show a complex biphasic time dependence which is particularly noticeable for the 100 and 200 PM curves.5 In the first several seconds, there is a rapid rate of oxidation which then slows. This is followed by a second phase during which the oxidation rate accelerates, reaching a maximum shortly before exhaustion of the HzOa. A reversible inactivation process which is consistent with an early accumulation of enzyme in the nonperoxidative path is suggested by these results (Loop 2) while the Hz02 concentration is comparatively high. This behavior can be interpreted as substrate-dependent inhibition producing a dead-end complex (under Results, see Computer Simulations). In further contrast to the data in Fig. 5, significant catalatic activity is clearly manifested for the higher concentrations of H202. The stoichiometry is depressed to only 51% of the theoretical for the 200 PM addition compared to 100% conversion for the 20 PM case. Figure 8b plots the final concentration of TOL. produced versus the initial H,02 concentration. The general trend shows 5 This may be seen more clearly by inclining and sighting along each trace.
the graph at an angle
52
MOORE,
MORONNE,
a.
AND
MEHLHORN
b.
20 ,A4 H,O,
Time (sec.) FIG. 8. TOLH oxidation vs HRP for low phenol. (a) The conditions for this experiment are as described in the legend to Fig. 5 except that 40 pM phenol is used instead. (b) Plot of the final TOL * vs the initial H,O, concentration. The dotted lines in (a) are theoretical fits using computer simulations of the model shown in Fig. 1 with allowance made for the catalatic effect of TOL . (see Computer Simulations under Results). The rate constant for the simulated curves are kl = 1.2 X 10’ Mm’ s-l, k2 = 2 X 10s Mm' SC', k3 = 1.2 X lo5 Me' s-l, k4 = 650 Mm' s-l, k, = 0.04 s-l, ks = 5 X lo4 M-' s-l, k7 = 500 M-' s-l, $ = 20 M-l s-l HRP-‘.
a relative loss of oxidizing equivalents for increased initial concentrations of HzOz. This result is in keeping with the model proposed by Kohler et al. (3), who found that the reconversion of CIII to native enzyme for LPO and TPO generated approximately 1 mol of O2 for every 2 mol of HaOz consumed. Presumably, this results from the release of O2 by CIII to give the ferrous enzyme (CIV), which then combines with H202 to give CR. However, as is discussed (under Computer Simulations), it appears that this mechanism is unable to account for the large amount of H202 decomposition observed under conditions of low phenol concentrations. Recent results (16) indicate that TOL * is able to promote catalatic activity in the presence of heme proteins and is able to account for the discrepancy. Effects of Phenol on Substrate-Dependent
Inhibition
The HzOz-dependent inhibition of the peroxidase enzyme activity is a function of the electron donor concentration [e.g., Fig. 5 vs Figs. 8a and 8b, this paper; and Ref. (7)]. Figure 9 shows the effects of increasing phenol concentration on the initial TOLH oxidation rates mediated by HRP and LPO. The rate of oxidation begins to saturate for both enzymes as the phenol concentration is raised. Phenol produces much the same increase in the LPOmediated rate of TOLH oxidation as it does for HRP. However, for the same unit concentration of enzyme (pyrogallol units) and the same phenol concentration, LPO appears to have about twice the activity of HRP. Interestingly, in contrast to the HRP results, biphasic kinetics were not observed with LPO even at low phenol concentrations (40 PM). This suggests that the rate of CIII formation for LPO is either slower than that for HRP or that the phenol enhancement of the peroxidase cycling is faster, producing less CIII and CIV.
Computer Simulations In order to validate the reaction cycle in Fig. 1, two notable features of the experimental data must be explained by the proposed model. First, the model must be able to generate the biphasic characteristics of the TOLH oxidation curves evident in samples containing comparatively low concentrations of phenol and large concentrations of HzOZ (Fig. 8). Second, the model must account for the decreasing conversion efficiency of HzOz equivalents to TOL * under the same conditions. Computer simulations using numerical solutions to Eqs. [l-3] indicate that biphasic kinetics of TOLH oxidation can be produced by the Kohler model. Figures 10a and lob show the predicted time-dependent concentrations of TOL a, HtOa, and enzyme components in a system with 40 PM phenol, 200 PM H202, and 2 U/ml HRP (same as in Fig. 8). Two cases are considered: (i) the basic Kohler model as depicted in Fig. 1 (solid lines) and (ii) a modified
--c) Q-----------------30
-I
Kl
.’
*’
.-
.-
.’
.*’
, ,’ P’ I’ I’ .’ t A.
,&----
_--- --
. o- LPO A- HRP
--*-------------------
FIG. 9. Initial TOLH oxidation rates vs phenol. Phenol stimulation of TOLH oxidation for HRP and LPO. Enzyme = 1 U/ml, 50 pM H,Oa, 2.5 mM TOLH, pH 7.2, at 22°C.
PEROXIDASE
ACTIVITY
BY ELECTRON
SPIN
53
RESONANCE
a.
200 f
HA 150
d.
b.
260
360
Time (sec.) FIG. 10. (a-d) Simulations based on the model in Fig. 1. (a) Predicted concentrations of HZ02 and TOL. in the case of 40 /.tM phenol (solid and dashed lines represent unmodified and modified models respectively); (b) corresponding enzyme concentrations. (c and d) The same quantities, but calculated for the case of 500 PM phenol. In each case the following rate constants were used: k, = 1.2 X lo7 Mm1 s-l, k2 = 2 X lo6 Me’ se’, k, = 1.1 X lo5 M ’ s I, k, = 250 M-’ s ‘, k, = 0.05 s-l, k, = 5 X lo4 M-’ s-i, k, = 500 M-’ s-i, k8 = 20 M-’ s-i HRP.
version which includes correction for TOL * -dependent catalatic activity (dashed lines). Both models demonstrate the characteristic kinetic behavior associated with substrate inhibition. The initial rates of TOLH oxidation are comparatively rapid but then decline. Thereafter, the oxidation rates accelerate, reaching a maximum shortly before exhaustion of the H202. As shown in Fig. lob, this behavior is reflected in the concentration of nonperoxidative enzyme (CIII + CIV) that at first rapidly increases coincident with the early inactivation of peroxidase activity. Subsequently, as HzOz is consumed, the concentrations of CIII and CIV decrease, gradually restoring activity. Although the biphasic kinetics are well represented, the unmodified model predicts a yield very close to the maximum value of 2 TOL . per H,O, even for 40 PM phenol. This is not consistent with the experimental results in Fig. 8 that indicate substantial catalatic action with loss of H,O,. According to the simulation only 5 j.rM of 200 PM H,O, is lost to catalatic activity (Loop 2); this is in contrast to experimental results showing 95 PM H,Oz destroyed (Fig. 8). The computer result is obtained despite the fact that the value of k4 used in the simulation was 10 times higher than previous literature values (4, 17). Even with the higher rate constant, cycling through Loop
2 is apparently too slow to account for the observed destruction of H,Oz. Recent experiments with heme proteins have shown that nitroxides can promote catalatic action (16). Nitroxides are not consumed during this process suggesting the following: peroxidase
2H202 + TOL * -
ks
2H,0+02+TOL..
(5)
Without attempting to specify the details of this process, oximetry data yield an estimate of 20 M-’ s-l per unit HRP for k8 under the experimental conditions used in the current studies. Including this reaction and the estimated value for k8 in the computer model produces the dotted line traces shown in Figs. 10a and lob. In this case, the loss of Hz02 is very close to the experimental result. The adequacy of the model is further illustrated by the dotted theoretical lines shown in Fig. Ba. Each of the theoretical curves was calculated based on a fit to the experimental data for the 200 PM H,O, trace. The data for each concentration of Hz02 agree reasonably well with the calculated curves, indicating self-consistency. In addition, the disappearance of biphasic behavior for the
54
MOORE,
MORONNE,
lowest concentrations of HzOz, as well as the near stoichiometric conversion of HaOz to TOL . , is also properly indicated. Figures 1Oc and 10d show the predicted rate of TOLH oxidation and the various enzyme forms when the electron donor concentration (phenol) is increased to 500 pM (as in Fig. 5). In this case, the TOLH oxidation rate is increased an order of magnitude (note the time scale), biphasic kinetics are no longer apparent, and the catalatic decomposition of Hz02 is greatly reduced. Again these data are consistent with the experimental results in Figs. 5 and 8. DISCUSSION
Measurement
of H202 and Peroxidase Activity
by ESR
We have shown that ESR methods can be used to measure trace activities of peroxidase enzymes (e.g., HRP activities as low as 0.003 U/ml pyrrogallol units). The assay is suitable for peroxidase enzymes that can be obtained free of reducing substrates. If this condition is fulfilled, then a substrate concentration-dependent stimulation of TOLH oxidation in the presence of hydrogen peroxide is sufficient to demonstrate the presence of peroxidase activity. This presumes that the oxidized substrates are capable of converting TOLH to its radical form [reduction potential = 0.3 V for the nearly identical TEMPOH, Ref. (IS)]. In this regard, the peroxidase oxidation products of biologically important substrates such as phenols and halides are potent TOLH oxidants. Further, the rates of reaction with TOLH are sufficiently fast that production of TOL . is an accurate measure of the rate of production of the oxidized intermediate. The fact that the peroxidase substrate is regenerated upon reaction with TOLH means that its concentration is effectively clamped, helping to maintain a steady rate of TOLH oxidation and enzyme cycling. This is a particular analytical advantage since the evaluation of activity is not limited to precise extrapolations of initial rates which are often complicated by the presence of noise and other artifacts such as mixing transients. Another advantage is that the ESR approach is sparing of biological materials. For example, with a typical 20-4 ESR sample the activity of as little as 6 pmol of enzyme can be measured. This sensitivity can be increased by an order of magnitude using a loop gap resonator which requires a sample size less than 2 ~1 (19). The efficacy of the ESR assay suggests its utility in systems where sensitivity and response time are important, such as samples with either very low or high activity. In the case of the latter, reaction rates on the millisecond time scale can be measured with ESR stopped-flow. For samples with low activity the accumulation of submicromolar concentrations of the stable radical product TOL . can be readily followed free of optical interference. HzOz and organic hydroperoxides capable of forming CI
AND
MEHLHORN
can be also assayed with peroxidase and an excess of an electron donor such as phenol so that peroxide is stoichiometrically converted to TOL * (Fig. 5). In addition, HZOz produced enzymatically can be quantitated using high peroxidase and electron donor concentrations. In this instance the rate of TOL . production will be determined by the rate of HzOz generation (e.g., Fig. 7). The basic constraint of the ESR assay and peroxidase assays in general is that conditions be chosen to minimize enzyme inactivation by conversion either to inactive but reversible forms such as CIII and CIV or to fully degraded enzyme. Inactivation is promoted by prolonged exposure to high concentrations of H202. As a consequence, peroxidase activity is best measured with low levels of Hz02 ( 0.5 mM produce simple kinetics and give nearly theoretical conversion of HzOz to TOL . . There have been a number of recent studies on the formation of CIII by high levels of HzOz and the reconversion of CIII to native enzyme (3, 4, 7, 9, 14, 17). In regard to the ESR assay, pertinent mechanisms identified for Compound III formation include (i) the direct reaction of HzOZ with CII (3, 4) and (ii) addition of superoxide anion to ferriperoxidase (4, 22, 23). In the ESR system, the former reaction seems highly probable and is consistent with increasing inhibition of TOLH oxidation with increasing HZOz. However, the direct participation of superoxide seems unlikely. It has been determined that reduced nitroxides similar to TOLH scavenge superoxide, producing TOL . and HzOz. Rosen et al. (24) determined a rate constant of -lo3 M-l s--l for superoxide and TEMPOH, a close analog of TOLH. Thus, the relatively high concentration of TOLH (2.5 mM) used in these experiments argues against the participation of freely diffusible superoxide. In addition, nitroxides such as TOL . rapidly form superoxide adducts that are potent oxidants of thiols and probably TOLH. The rate constant for adduct formation is estimated to be 3 X 10” M-’ s-’ (25). As a consequence, TOL * produced during TOLH oxidation is most likely to increase superoxide scavenging. Although it appears unlikely that superoxide contributes to CIII formation in the ESR-TOLH system, analyses of experiments using low phenol (40 PM, Fig. 8) indicate that the rate of CIII formation is enhanced compared to spectrophotometric measurements done in the absence of electron donors (4, 17). To fit data such as those in Fig. 8, it. was found that k, (Fig. 1) needed to be 5-20 times higher than the previous values of 20-25 M ’ so) (4, 17). Simulations using the slower literature value did not produce a sufficient rate of peroxidase inactivation to account for either the initial inactivation phase or the subsequent recovery of activity; in other words, too little enzyme is directed through Loop 2. This result suggests that phenol, TOLH, or TOL. is able to directly or indirectly facilitate the conversion of CII to CIII. In the case of phenol, Ma and Rokita (14) determined that the rate of irreversible peroxidase inactivation of HRP by Hz02 was increased an order of magnitude when phenol was included in the reaction mixture. Since it has been argued that CIII is an intermediate that leads to irreversible inactivation (3), this may indicate that phenol or its radical products facilitate formation of CIII. The
SPIN
55
RESONANCE
involvement of phenoxyl radicals in the ESR system seems unlikely since the steady-state concentration is expected to be very low given the rapid scavenging provided by TOLH. The possibility that TOLH is able to promote CIII formation also seems remote. Experiments comparing 0.1, 0.5, and 2.5 mM TOLH yielded the same overall rates of TOLH oxidation, suggesting that formation of CIII is unaffected by TOLH. As discussed earlier, TOL * promotes the catalatic activity of heme proteins including HRP. The exact mechanism remains to be elucidated. However, one possible scenario involves the oxidation of TOL . to the immonium oxene cation [TOL+, Ref. (26)] by CI, CII, or phenoxyl. The oxene compound may oxidize H202 to give water and oxygen, thus acting as a catalase, or might give an adduct capable of converting native enzyme to CIII. When the initial TOL . concentration was increased from 90 to 450 PM in a mixture consisting of 1 U/ml HRP, 100 PM HzOz, and 0.5 mM phenol, a 20% drop in conversion efficiency and peroxidase activity (oxidation rate) resulted. The fact that the activity declined indicates that high TOL . may promote CIII formation. ACKNOWLEDGMENTS We thank Dr. Judith Klinman of the Department of Chemistry, UC Berkeley, for helpful discussions. This work was supported by NIH Grant AG 04818, the Cigarette and Tobacco Surtax Fund of the State of California through the Tobacco-Related Disease Research Program of the University of California, and Office of Energy Research, through the Department of Energy under Contract DE-AC03-76SF00098.
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12. Moronne, M. M., Mehlhorn, R. J., Miller, M. P., Ackerson, L. C., and Macey, R. I. (1990) J. Memb. Biol. 115, 31-40. 13. Prolla, T. A., and Mehlhorn, R. J. (1990) Free Rd. Res. Commun. 9,135-146. 14. Ma, X., and Rokita, S. E. (1988) Biochem. Biophys. Res. Commun. 157(l), 170-175. 15. Schuler, R. H. (1977) Radiat. Res. 69, 417-433. 16. Mehlhorn, R., and Swanson, C. E. (1992) Free Rd. Res. Commun., in press. 1’7. Adediran, S. A., and Lambeir, A. (1989) Eur. J. Biochem. 186,571I Fil6. 18. Rosen, G. M., Rauckman,
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20. Moore, K. L., (1990) PhD dissertation, Berkeley. 21. Jenzer, H., and Kohler,
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Radicals (Ulrich,
H., Ed.),
