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pH kinetic studies of the N-demethylation of N,N-dimethylaniline catalyzed by chloroperoxidase
ARCHIVES OF BIOCHEMISTRY Vol. 233, No. 2, September,
AND BIOPHYSICS pp. 315-321, 1984
pH Kinetic Studies of the N-Demethylation of N,N-Dimethylaniline Catalyzed by Chloroperoxidase’ GREGORY Departments
of Pathology, Northwestern Received
L. KEDDERIS2
AND
PAUL
F. HOLLENBERG3
and Molecular Biology and Biochemistry, and the Cancer University Medical School, Chicago, Illinois 60611 October
31, 1983, and in revised
form
April
Center,
25, 1984
The effect of pH on the kinetic parameters for the chloroperoxidase-catalyzed Ndemethylation of N,N-dimethylaniline supported by ethyl hydroperoxide was investigated from pH 3.0 to 7.0. Chloroperoxidase was found to be stable throughout the pH range studied. Initial rate conditions were determined throughout the pH range. The I’,,,,, for the demethylation reaction exhibited a pH optimum at approximately 4.5. The K, for N,N-dimethylaniline increased with decreasing pH, while the K, for ethyl hydroperoxide varied in a manner paralleling V,,,. Comparison of the V,,,/K, values for N,N-dimethylaniline and ethyl hydroperoxide indicated that the interaction of N,N-dimethylaniline with chloroperoxidase compound I was rate-limiting below pH 4.5, while compound I formation was rate-limiting above pH 4.5. The log of the V,,,/ K, for ethyl hydroperoxide was independent of pH, indicating that chloroperoxidase compound I formation is not affected by ionizations in this pH range. The plot of the log of the V,,,/K, for N,N-dimethylaniline versus pH indicated an ionization on compound I with a pK of approximately 6.8. The plot of the log of the V,,, versus pH indicated an ionization on the compound I-N,N-dimethylaniline complex, with a pK of approximately 3.1. The results show that chloroperoxidase can demethylate both the protonated and neutral forms of N,N-dimethylaniline (pK approximately 5.0), suggesting that hydrophobic binding of the arylamine substrate is more important in catalysis than ionic bonding of the amine moiety. For optimal catalysis, a residue in the chloroperoxidase compound I-N,N-dimethylaniline complex with a pK of approximately 3.1 must be deprotonated, while a residue in compound I with a pK of approximately 6.8 must be protonated.
Chloroperoxidase (chloride:hydrogen peroxide oxidoreductase; EC 1.11.1.10) is a
monomeric hemeprotein isolated from the mold Caldariomyces fumago. Chloroperoxidase (CPO)4 utilizes peroxy compounds to catalyze a variety of different reactions. In the presence of halide anions (Cl-, Br-, or II, but not F-), this enzyme catalyzes the peroxidative formation of a carbon-halogen bond with several P-diketones and aromatic compounds (1). CPO also catalyzes the classical peroxidative oxidation of a wide variety of organic molecules, and can decompose hydroperoxides and peracids to
i This research was supported by Grant CA-16954 from the National Institute of Health. Portions of this work were presented at a meeting of the American Society of Biological Chemists at St. Louis, MO., June 1981. *The data in this paper are taken from a thesis submitted by G.L.K. in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biochemistry in the Graduate School of Northwestern University. Present address: Chemical Industry Institute of Toxicology, P.O. Box 12137, Research Triangle Park, N. C. 27709. ‘To whom reprint requests should be sent.
’ Abbreviations N,N-dimethylaniline; 315
used: CPO, chloroperoxidase; DMA, EtOOH, ethyl hydroperoxide. 0003-9861/84
$3.00
Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved.
316
KEDDERIS
AND
give molecular oxygen (2). The enzyme also catalyzes the oxidation of aniline and its derivatives to nitrosobenzene (3). Additionally, CPO catalyzes the N-demethylation of secondary and tertiary Nmethylarylamines (4) according to ROOH + R’R”N-CHS R’R”NH
+ ROH + HCHO,
[l]
where ROOH is the hydroperoxide and RR’N-CHS is the N-methylarylamine substrate. A steady-state kinetic analysis of the CPO-catalyzed N-demethylation of N,N-dimethylaniline (DMA) indicated that the reaction proceeds by a Ping-Pong mechanism involving the substituted enzyme intermediate compound I (5). In this mechanism, EtOOH reacts with the peroxidase to form the oxidized intermediate compound I with the concomitant release of ethanol. DMA then binds to compound I and is oxidized, resulting in the formation of N-methylaniline and formaldehyde and the regeneration of the native enzyme. We have previously reported that the CPOcatalyzed demethylation of DMA is a pH-dependent reaction with an apparent pH optimum at approximately 6.0 (4). In order to investigate the roles of enzyme and substrate ionizations in catalysis, we have studied the effect of pH on the kinetic parameters of the demethylation reaction. The results of this study demonstrate that CPO can efficiently demethylate both the protonated and neutral forms of DMA, and that the rate of the demethylation reaction is controlled by two ionizations on the enzyme. EXPERIMENTAL
PROCEDURES
Elzzyne preparation Chloroperoxidase was isolated and purified from C. fumago using the methods reported previously (1). The enzyme preparations used in these studies had specific activities greater than 2000 units/mg protein in the standard assay for the chlorination of monochlorodimedone (l), and exhibited Am/Also ratios greater than 1.40, indicating that the enzyme preparations were at least 95% pure. Protein concentrations were determined by the method of Lowry et al. (6), using bovine serum albumin as the standard. Materiuk N,N-Dimethylaniline was obtained from Aldrich Chemical Company, and was redistilled before
HOLLENBERG use. Ethyl hydroperoxide was obtained from Polysciences Inc., and its concentration was determined by iodometric titration (7). All other materials were reagent grade, obtained from commercial sources. Preparation of In&bra Phosphate buffers were prepared by dissolving the appropriate amount of sodium phosphate (monobasic) in glass-distilled water, and titrating with concentrated potassium hydroxide solution. Phosphate buffers were prepared at pH values below pH 4.5 by diluting the appropriate amount of phosphoric acid in glass-distilled water, and titrating with concentrated potassium hydroxide solution. All pH measurements were made on a Corning Model 110 digital pH meter equipped with a Sensorex combination electrode. Assay for demethylation The N-demethylase activity of CPO was determined by measuring the amount of formaldehyde formed using a modification of the procedure of Nash (8) as previously described (4). In the pH range from 4.5 to 7.0, DMA was added to the reaction mixtures (3 ml final volume) in less than 15 ~1 acetone. This concentration of acetone has no effect on the demethylation reaction under these conditions. In the pH range from 3.0 to 4.5, DMA was added to the reaction mixtures in phosphate buffer titrated to the appropriate pH with potassium hydroxide solution. The reactions were initiated by the addition of CPO, incubated at 25°C for the times indicated, and terminated by the addition of 0.75 ml 60% trichloroacetic acid. Some of the reaction mixtures were then extracted with 5 ml ethyl acetate to remove the violet color formed during the course of the reaction (4). Since the ethyl acetate extraction step did not significantly change the slope of formaldehyde standard curves, incubations where no violet color was formed’were not extracted. At pH 3.5 and 3.0, the terminated reaction mixtures were subsequently neutralized with potassium hydroxide and extracted again with ethyl acetate to remove excess DMA. The high concentrations of DMA used at these pH values interfered with the absorbance readings if the DMA was not extracted before addition of the Nash reagent. A l-ml aliquot of the aqueous phase was mixed with 0.5 ml Nash reagent (30 g ammonium acetate and 0.4 ml 2,4-pentanedione per 50-ml volume) and incubated at 25°C for 45 min. The absorbance of the resulting conjugate was measured at 421 nm on a Gilford 2400-S uv-visible spectrophotometer. Initial rate conditions for the demethylation reaction (linearity with time and enzyme concentration) were determined at each pH value used in these studies. The reaction was linear with time for at least 10 min at pH values from 3.5 to 6.0, and for at least 5 min at the pH values 3.0, 6.5, and 7.0. The apparent kinetic parameters for the demethylation reaction were determined from double-reciprocal plots of data obtained at concentrations of the fixed substrate which
pH
STUDIES
ON
N-DEMETHYLATION
were approximately three times its K,,,, and linear plots were obtained in all cases. The results of a typical experiment are shown in Fig. 1. Higher concentrations of the fixed substrate were not possible at the extremes of the pH range. At pH 6.5 and 7.0, marked substrate inhibition by DMA was observed at higher concentrations, while at pH 3.0, solutions of DMA greater than 50 mM were not feasible due to insolubility. All kinetic determinations were carried out using concentrations of the varied substrate that gave an equal distribution of points around the apparent K,,, obtained in double-reciprocal plots of the data. At least seven different concentrations of the varied substrate were used in each experiment. All experiments were performed at least twice, with each point done in duplicate.
RESULTS
AND
DISCUSSION
CPO was found to be stable throughout the pH range under investigation. The enzyme was preincubated for 15 min at 25°C in buffers spanning the pH range, and then assayed for demethylation activity at pH 6.0. The same amounts of formaldehyde were formed from each sample of CPO preincubated at the various pH values, indicating that the pH-dependence of the rate of demethylation was not due to enzyme inactivation at the extremes of pH. The change in the rate of the demethylation reaction in the pH range from 3.0 to 7.0 may be due to ionizations of the substrates or of groups on the enzyme. EtOOH,
CHLOROPEROXIDASE
0.2 [N, N-Dimethylaniline]-’
0.4
0.6 (m&l
317
with a pK of 11.8 (9), does not ionize over this pH range, while DMA, with a pK around 5.0 under a wide variety of conditions (lo-13), will ionize in the pH range studied. In order to elucidate the roles of enzyme and substrate ionizations, the effect of pH on the kinetic parameters of the demethylation reaction was determined throughout the pH range of 3.0 to 7.0, and the results were analyzed as described by Tipton and Dixon (14). Initial rate and inhibition studies of the CPO-catalyzed N-demethylation of DMA indicated that the reaction proceeds by a Ping-Pong kinetic mechanism (5). Thus, for the demethylation reaction, the I’,,,,,/ Km for each substrate (EtOOH or DMA) is the second-order rate constant for the binding and reaction of each substrate with the appropriate enzyme species (native CPO or compound I), and the pH dependence of the V,,,,,/K, will indicate kinetically important ionizations of the free substrate or enzyme species (14). Similarly, the pH dependence of the V,, will indicate kinetically important ionizations of the ES complex (14), since the I’,,,,, is the rate of the reaction at saturating concentrations of both substrates. For the demethylation reaction, the pH dependence of the V,,, will reflect ionizations on whichever ES complex (CPO-EtOOH or compound IDMA) is rate-limiting within the pH range. 8
8-
- 0.2
BY
-I
0.8 [Ethyl
Hydroperoxidel-‘(mM)-’
FIG. 1. Double-reciprocal plots of the initial rate of the demethylation of DMA at pH 4.5. The 3-ml reaction mixtures contained 0.5 M sodium potassium phosphate buffer, pH 4.5, and 0.29 yg CPO. (A) The concentration of DMA was varied as indicated in the presence of 5.36 mM EtOOH. (B) The concentration of EtOOH was varied in the presence of 5.26 mM DMA. The reactions were initiated by addition of the enzyme, and incubated for 10 min at 25’C. Formaldehyde formation was determined by the Nash assay as described under Experimental Procedures.
318
KEDDERIS
AND
The apparent kinetic parameters for the demethylation reaction as a function of pH are shown in Table I. The V,,, for the demethylation reaction exhibited a pH optimum of approximately 4.5. This is similar to the pH optima observed for the CPOcatalyzed evolution of oxygen from hydrogen peroxide (2) and for the N-oxidation of pchloroaniline (15). As shown in Table I, the Km for DMA decreased with increasing pH while the K, for EtOOH varied with pH in a manner paralleling V,,. The pH optimum of the V,,, for the demethylation reaction (pH 4.5) is different from the pH optimum of 6.0 previously reported (4). In the previous study (4), substrate concentrations which were saturating at pH 6.0 (0.53 mM DMA and 1.66 mM EtOOH) were used for the entire pH range, and the assumption was made that the initial velocities measured as a function of pH were proportional to the maximal velocities (14). However, it is apparent from the data in Table I that the decrease in the rate of the demethylation reaction with decreasing pH under the conditions used previously (4)
TABLE
I
EFFECT OF pH ON THE KINETIC PARAMETERS FOR CHLOROPEROXIDASE-CATALYZED DEMETH~ATION OF N,N-DIMETHYLANILINE’
K,,, DMA
PH 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0
@M) 23.27 13.57 5.36 3.05 0.809 0.382 0.088 0.061 0.034
f f f + It k * f +
K,,, EtOOH
(mM) 0.76 0.72 0.73 0.10 0.005 0.034 0.007 0.005 0.004
1.34 1.66 2.20 2.53 2.01 1.77 0.889 0.855 0.498
+ * f 5 I+ f * f k
0.01 0.03 0.10 0.09 0.23 0.05 0.022 0.021 0.063
V,, (nmol formaldehyde min-’ pg CPO-‘)
56.4 104.0 127.9 160.4 141.5 132.8 81.7 82.7 57.7
k f f + + k f + +-
6.6 13.6 3.3 5.6 9.0 8.1 12.8 5.4 6.0
“The reaction mixtures in sodium potassium phosphate buffer (0.5 M), pH as indicated, were initiated by addition of chloroperoxidase, incubated at 25°C for the times indicated under Results, terminated by addition of 60% trichloroacetic acid, and assayed for formaldehyde formation as described under Experimental Procedures. The chloroperoxidase concentrations were 0.895 ~3 at pH 3.0, 0.29 ~3 at pH 3.5 to 6.5, and 0.45 pg at pH 7.0. The data are presented as the mean values plus or minus standard errors from at least two determinations of K, and at least four determinations of V,,.
HOLLENBERG
was due to the use of subsaturating concentrations of DMA and EtOOH at pH values less than 6.0 rather than due to a ratecontrolling ionization. The data in Table I also indicate that CPO can efficiently demethylate both the protonated and neutral forms of DMA, suggesting that, under these conditions of ionic strength, hydrophobic binding of the DMA phenyl ring is more important in catalytic binding than ionic binding of the amine moiety. Comparison of the Vm,,/Km values for DMA and EtOOH (shown in Table II) indicates that the interaction of DMA with compound I is rate-limiting below pH 4.5, while compound I formation is rate-limiting above pH 4.5. Therefore, the pH dependence of the V,, for the demethylation reaction will reflect ionizations on the compound I-DMA complex at pH values below 4.5 and ionizations on the CPOEtOOH complex at pH values above 4.5. The V,,,,,/K, for DMA is approximately 700 times greater at pH 7.0 than at pH 3.0. In contrast, the V,,,/K, for EtOOH changes by less than a factor of three as the pH increases from 3.0 to 7.0. The kinetic constants in Table I were analyzed as described by Tipton and Dixon (14) in order to identify kinetically important ionizations of the free substrates and enzyme species. As shown in Fig. 2, the plot of the log of V,,,/K, versus pH was essentially independent of pH for EtOOH. Since the V,,,,,/K, for EtOOH represents the second-order rate constant for the reaction of EtOOH with CPO to form compound I, these results indicate that compound I formation was not affected by any ionizations in the pH range studied. The curvature in the plot at the extremes of pH is not great enough to indicate any ionizations, and is probably due to rate-controlling ionizations outside of the pH range studied. The V,,/K, for DMA represents the second-order rate constant for the reaction of DMA with CPO compound I. The plot of the log of the V,,,/K, for DMA (Fig. 3) increased with increasing pH in a linear fashion (slope = 1) through pH 6.0 and then leveled off. Extrapolation of the straight-line portions of the plot indicated
pH
STUDIES TABLE
ON
N-DEMETHYLATION
BY
319
CHLOROPEROXIDASE
II
EFFECT OF pH ON THE V/K VALUES FOR CHLOROPEROXIDASE-CATALYZED DEMETHYLATION OF N,N-DIMETHYLANILINE” V,,,,,/K,,, (nmol formaldehyde min-’ CPO-’ ITIM-‘)
fig
IO
PH
3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5
1360
7.0
1700
23.9
52.6 175 348
from
/ PH
FIG. 3. Dependence of the log of the V,,,,/K,,, for N,N-dimethylaniline upon pH. The VmI,../K,,, was calculated from the data in Table I. The standard errors in the V,,,,,/K,,, were determined by the propagation of error in the kinetic parameters.
92.0 96.7
928
a Calculated
05
42.1 62.7 58.1 63.4 70.4 75.0
2.42 7.66
116
the data
in Table
I.
a pK of approximately 6.8 on compound I or DMA. Since the pK of DMA is around 5.0 (lo-13), this must be an ionization of compound I. Thus, the ionization of DMA, although occurring in this pH range, is not important for catalysis since it does not show up in the kinetic plots. If the increase in the Km for DMA with decreasing pH (Table I) was due to its protonation to a form which could not be demethylated by the enzyme, then redefinition of the Km in terms of the concentration of the neutral species of DMA at a given pH should result
1,
I 30
1 40
I
1 50 PH
I
I 60
I
I 70
FIG. 2. Dependence of the log of the V,,/K, for ethyl hydroperoxide upon pH. The V,,/K,,, was calculated from the data in Table I. The standard errors in the V,‘,,/K,,, were determined by the propagation of error in the kinetic parameters.
in the pH independence of the Km. The Km can be redefined in terms of the concentration of the neutral species of DMA by the following modification of the Henderson-Hasselbach equation (16) Kk = K,/[l
+ antilog(pK
- pH)],
where Km is the apparent Michaelis constant at a given pH value, pK is the ionization constant of DMA (5.0), and K& is the Km in terms of the concentration of the neutral species of DMA. When this equation was used to redefine the apparent Km values for DMA in terms of the concentration of the neutral species of DMA, the redefined Km values were still pH dependent (not shown), further supporting the conclusion that the ionization of DMA is not catalytically important in the demethylation reaction. Extrapolation of linear tangents to the curved section of the plot of the log of V,,, versus pH (Fig. 4) at the lower limit of the pH range studied indicated a pK of approximately 3.1 on the compound I-DMA complex. The decrease in the log of V,,, at higher pH was not great enough to indicate a pK within the pH range studied, and is probably due to an ionization of the CPO-EtOOH complex with a pK greater than 7.0. This ionization is probably the
320
KEDDERIS
AND
244
2ot T---d 04I 30
1
I 40
1
1 50 PH
I
1 60
I
I 70
FIG. 4. Dependence of the log of the V,,,, for the demethylation reaction upon pH. The log of V,, was calculated from the data in Table I.
same group whose ionization in compound I was indicated in the plot of the log of Vm,,/Km for DMA versus pH (Fig. 3), its pK being shifted to a slightly higher value in the CPO-EtOOH complex. Similarly, the pK of approximately 3.1 indicated in the plot of the log of V,,,,, versus pH (Fig. 4) may be the same group responsible for the downward trend at low pH in the plot of the log of V,,,/K, for EtOOH versus pH (Fig. 2), its pK also being slightly higher in the compound I-DMA complex. This ionization may represent the transition between the acid and neutral forms of CPO, which has been reported to have a pK of approximately 3.5 (17). Hager and co-workers (18) have demonstrated that carbon monoxide binding by ferrous CPO is influenced by an ionizable group on the enzyme which exhibits a pK of approximately 5.5. The unprotonated form of the enzyme reacts with carbon monoxide much faster than the protonated form. Based on the pK and the AH“ (8 kcal mall’) for the ionization of the functional group affecting the binding of carbon monoxide, they suggest that this ionizable group is a histidine residue at the active site which is on the distal side close to the sixth axial ligand position of the heme iron atom. The studies described here on the CPO-catalyzed demethylation reaction do not provide any evidence for the involvement of a residue on the protein having a pK of 5.5. However, these studies involve the ferric as well as higher oxidation states
HOLLENBERG
of the heme iron, whereas in the carbon monoxide binding studies only the ferrous form could be investigated. Since variations in the electronic environment of amino acid residues in proteins may result in shifts in pK values of as much as 2-3 units (14), the ionizable group having a pK of approximately 6.8 which we observed in compound I may be the same imidazole histidine which has been shown by Hager and co-workers to play a role in the binding of carbon monoxide to ferrous CPO. However, the identities of the ionizable groups cannot be determined from the kinetic data presented here. The results of the pH kinetic analysis of the CPO-catalyzed demethylation reaction indicate that for optimal catalysis, a residue on the compound I-DMA complex with a pK of approximately 3.1 must be deprotonated while a residue on compound I with a pK of approximately 6.8 must be protonated. Marked substrate inhibition of the demethylation reaction by DMA was observed above pH 6.0 (data not shown), suggesting that deprotonation of the residue with a pK around 6.8 results in a higher affinity of DMA for an inhibitory site on the enzyme. The deprotonation of the group with a pK around 3.1 and the protonation of the group with a pK around 6.8 may be necessary to achieve the optimal conformation of the active site for catalysis. Over the pH range from 3.5 to 6.0, the two residues are mostly in their optimal ionization states for catalysis. At the extremes of the pH range, the active site may be in less optimal conformations, resulting in a slower rate of demethylation. ACKNOWLEDGMENTS We thank Professor Lowell P. Hager for supplying the purified chloroperoxidase and for his helpful suggestions and encouragement. We thank Ms. Nancy Starks for her excellent assistance in the preparation of this manuscript.
REFERENCES 1. MORRIS, D. R., AND HAGER, L. P. (1966). J. Biol. Chmz. 241, 1763-1’768. 2. THOMAS, J. A., MORRIS, D. R., AND HAGER, L. P. (1970) J. Biol. Chem 245, 3129-3134.
pH
STUDIES
ON
N-DEMETHYLATION
3. CORBEIT, M. D., BADEN, D. G., AND CHIPKO, B. R. (1979) B&n-g. Chm 8, 91-95. 4. KEDDF.RIS, G. L., KOOP, D. R., AND HOLLENBERG, P. F. (1980) J. Biol Chem. 255, 101’74-10182. 5. KEDDERIS, G. L., AND HOLLENBERG, P. F. (1983) J. BioL Chcm 258, 12413-12419. 6. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol Chem 193, 265-2’75. 7. SILBERT, L. S., AND SWERN, D. (1958) Ad Chem. 30,385-387. 8. NASH, T. (1953) Biochem J. 55,416-421. 9. EVERETT, A. J., AND MINKOFF, G. J. (1953) Trans. Faraday Sot. 49,410-414. 10. GUTBENZAHL, B., AND GRUNWALD, E. (1953) J. Amer. Chem. Sot 75, 559-565. 11. BACARELLA, A. L., GRUNWALD, E., MARSHALL, H. P., AND PURLEE, E. L. (1955) J. Org. Chem 20, 747-762.
BY
321
CHLOROPEROXIDASE
12. FICKLING, M. M., FISCHER, PACKER, J., AND VAUGHN, Chem Sot. 81.4226-4230. 13. CHO, A. K., AND MIWA, LXsp. 2, 477-483.
A., MANN, B. R., J. (1959) J. Amer.
G. T. (1973)
Drug
Met&
14. Tipton, K. F., and Dixon, H. B. F. (1979) in Methods in Enzymology (Purich, D. L., ed.), Vol. 63, pp. 183-234, Academic Press, New York. 15. CORBEIT, M. D., CHIPKO, A. 0. (1980) Biochem
B. R., AND BATCHELOR, J. 187, 893-903.
16. MCEWEN, C. M., JR., SASAKI, G., AND LENZ, W. R., JR. (1968) J. Biol Chew 243, 5217-5225. 17. THOMAS, (1970)
J. A., MORRIS, J. Bid Chem.
D. R., AND HAGER, 245, 3135-3142.
L. P.
18. CAMPBELL, B. A., JR., ARAISO, T., REINISCH, L., YUE, K. T., AND HAGER, L. P. (1982) Biochemistry 21, 4343-4349.
AND BIOPHYSICS pp. 315-321, 1984
pH Kinetic Studies of the N-Demethylation of N,N-Dimethylaniline Catalyzed by Chloroperoxidase’ GREGORY Departments
of Pathology, Northwestern Received
L. KEDDERIS2
AND
PAUL
F. HOLLENBERG3
and Molecular Biology and Biochemistry, and the Cancer University Medical School, Chicago, Illinois 60611 October
31, 1983, and in revised
form
April
Center,
25, 1984
The effect of pH on the kinetic parameters for the chloroperoxidase-catalyzed Ndemethylation of N,N-dimethylaniline supported by ethyl hydroperoxide was investigated from pH 3.0 to 7.0. Chloroperoxidase was found to be stable throughout the pH range studied. Initial rate conditions were determined throughout the pH range. The I’,,,,, for the demethylation reaction exhibited a pH optimum at approximately 4.5. The K, for N,N-dimethylaniline increased with decreasing pH, while the K, for ethyl hydroperoxide varied in a manner paralleling V,,,. Comparison of the V,,,/K, values for N,N-dimethylaniline and ethyl hydroperoxide indicated that the interaction of N,N-dimethylaniline with chloroperoxidase compound I was rate-limiting below pH 4.5, while compound I formation was rate-limiting above pH 4.5. The log of the V,,,/ K, for ethyl hydroperoxide was independent of pH, indicating that chloroperoxidase compound I formation is not affected by ionizations in this pH range. The plot of the log of the V,,,/K, for N,N-dimethylaniline versus pH indicated an ionization on compound I with a pK of approximately 6.8. The plot of the log of the V,,, versus pH indicated an ionization on the compound I-N,N-dimethylaniline complex, with a pK of approximately 3.1. The results show that chloroperoxidase can demethylate both the protonated and neutral forms of N,N-dimethylaniline (pK approximately 5.0), suggesting that hydrophobic binding of the arylamine substrate is more important in catalysis than ionic bonding of the amine moiety. For optimal catalysis, a residue in the chloroperoxidase compound I-N,N-dimethylaniline complex with a pK of approximately 3.1 must be deprotonated, while a residue in compound I with a pK of approximately 6.8 must be protonated.
Chloroperoxidase (chloride:hydrogen peroxide oxidoreductase; EC 1.11.1.10) is a
monomeric hemeprotein isolated from the mold Caldariomyces fumago. Chloroperoxidase (CPO)4 utilizes peroxy compounds to catalyze a variety of different reactions. In the presence of halide anions (Cl-, Br-, or II, but not F-), this enzyme catalyzes the peroxidative formation of a carbon-halogen bond with several P-diketones and aromatic compounds (1). CPO also catalyzes the classical peroxidative oxidation of a wide variety of organic molecules, and can decompose hydroperoxides and peracids to
i This research was supported by Grant CA-16954 from the National Institute of Health. Portions of this work were presented at a meeting of the American Society of Biological Chemists at St. Louis, MO., June 1981. *The data in this paper are taken from a thesis submitted by G.L.K. in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Biochemistry in the Graduate School of Northwestern University. Present address: Chemical Industry Institute of Toxicology, P.O. Box 12137, Research Triangle Park, N. C. 27709. ‘To whom reprint requests should be sent.
’ Abbreviations N,N-dimethylaniline; 315
used: CPO, chloroperoxidase; DMA, EtOOH, ethyl hydroperoxide. 0003-9861/84
$3.00
Copyright 0 1984 by Academic Press, Inc. All rights of reproduction in any form reserved.
316
KEDDERIS
AND
give molecular oxygen (2). The enzyme also catalyzes the oxidation of aniline and its derivatives to nitrosobenzene (3). Additionally, CPO catalyzes the N-demethylation of secondary and tertiary Nmethylarylamines (4) according to ROOH + R’R”N-CHS R’R”NH
+ ROH + HCHO,
[l]
where ROOH is the hydroperoxide and RR’N-CHS is the N-methylarylamine substrate. A steady-state kinetic analysis of the CPO-catalyzed N-demethylation of N,N-dimethylaniline (DMA) indicated that the reaction proceeds by a Ping-Pong mechanism involving the substituted enzyme intermediate compound I (5). In this mechanism, EtOOH reacts with the peroxidase to form the oxidized intermediate compound I with the concomitant release of ethanol. DMA then binds to compound I and is oxidized, resulting in the formation of N-methylaniline and formaldehyde and the regeneration of the native enzyme. We have previously reported that the CPOcatalyzed demethylation of DMA is a pH-dependent reaction with an apparent pH optimum at approximately 6.0 (4). In order to investigate the roles of enzyme and substrate ionizations in catalysis, we have studied the effect of pH on the kinetic parameters of the demethylation reaction. The results of this study demonstrate that CPO can efficiently demethylate both the protonated and neutral forms of DMA, and that the rate of the demethylation reaction is controlled by two ionizations on the enzyme. EXPERIMENTAL
PROCEDURES
Elzzyne preparation Chloroperoxidase was isolated and purified from C. fumago using the methods reported previously (1). The enzyme preparations used in these studies had specific activities greater than 2000 units/mg protein in the standard assay for the chlorination of monochlorodimedone (l), and exhibited Am/Also ratios greater than 1.40, indicating that the enzyme preparations were at least 95% pure. Protein concentrations were determined by the method of Lowry et al. (6), using bovine serum albumin as the standard. Materiuk N,N-Dimethylaniline was obtained from Aldrich Chemical Company, and was redistilled before
HOLLENBERG use. Ethyl hydroperoxide was obtained from Polysciences Inc., and its concentration was determined by iodometric titration (7). All other materials were reagent grade, obtained from commercial sources. Preparation of In&bra Phosphate buffers were prepared by dissolving the appropriate amount of sodium phosphate (monobasic) in glass-distilled water, and titrating with concentrated potassium hydroxide solution. Phosphate buffers were prepared at pH values below pH 4.5 by diluting the appropriate amount of phosphoric acid in glass-distilled water, and titrating with concentrated potassium hydroxide solution. All pH measurements were made on a Corning Model 110 digital pH meter equipped with a Sensorex combination electrode. Assay for demethylation The N-demethylase activity of CPO was determined by measuring the amount of formaldehyde formed using a modification of the procedure of Nash (8) as previously described (4). In the pH range from 4.5 to 7.0, DMA was added to the reaction mixtures (3 ml final volume) in less than 15 ~1 acetone. This concentration of acetone has no effect on the demethylation reaction under these conditions. In the pH range from 3.0 to 4.5, DMA was added to the reaction mixtures in phosphate buffer titrated to the appropriate pH with potassium hydroxide solution. The reactions were initiated by the addition of CPO, incubated at 25°C for the times indicated, and terminated by the addition of 0.75 ml 60% trichloroacetic acid. Some of the reaction mixtures were then extracted with 5 ml ethyl acetate to remove the violet color formed during the course of the reaction (4). Since the ethyl acetate extraction step did not significantly change the slope of formaldehyde standard curves, incubations where no violet color was formed’were not extracted. At pH 3.5 and 3.0, the terminated reaction mixtures were subsequently neutralized with potassium hydroxide and extracted again with ethyl acetate to remove excess DMA. The high concentrations of DMA used at these pH values interfered with the absorbance readings if the DMA was not extracted before addition of the Nash reagent. A l-ml aliquot of the aqueous phase was mixed with 0.5 ml Nash reagent (30 g ammonium acetate and 0.4 ml 2,4-pentanedione per 50-ml volume) and incubated at 25°C for 45 min. The absorbance of the resulting conjugate was measured at 421 nm on a Gilford 2400-S uv-visible spectrophotometer. Initial rate conditions for the demethylation reaction (linearity with time and enzyme concentration) were determined at each pH value used in these studies. The reaction was linear with time for at least 10 min at pH values from 3.5 to 6.0, and for at least 5 min at the pH values 3.0, 6.5, and 7.0. The apparent kinetic parameters for the demethylation reaction were determined from double-reciprocal plots of data obtained at concentrations of the fixed substrate which
pH
STUDIES
ON
N-DEMETHYLATION
were approximately three times its K,,,, and linear plots were obtained in all cases. The results of a typical experiment are shown in Fig. 1. Higher concentrations of the fixed substrate were not possible at the extremes of the pH range. At pH 6.5 and 7.0, marked substrate inhibition by DMA was observed at higher concentrations, while at pH 3.0, solutions of DMA greater than 50 mM were not feasible due to insolubility. All kinetic determinations were carried out using concentrations of the varied substrate that gave an equal distribution of points around the apparent K,,, obtained in double-reciprocal plots of the data. At least seven different concentrations of the varied substrate were used in each experiment. All experiments were performed at least twice, with each point done in duplicate.
RESULTS
AND
DISCUSSION
CPO was found to be stable throughout the pH range under investigation. The enzyme was preincubated for 15 min at 25°C in buffers spanning the pH range, and then assayed for demethylation activity at pH 6.0. The same amounts of formaldehyde were formed from each sample of CPO preincubated at the various pH values, indicating that the pH-dependence of the rate of demethylation was not due to enzyme inactivation at the extremes of pH. The change in the rate of the demethylation reaction in the pH range from 3.0 to 7.0 may be due to ionizations of the substrates or of groups on the enzyme. EtOOH,
CHLOROPEROXIDASE
0.2 [N, N-Dimethylaniline]-’
0.4
0.6 (m&l
317
with a pK of 11.8 (9), does not ionize over this pH range, while DMA, with a pK around 5.0 under a wide variety of conditions (lo-13), will ionize in the pH range studied. In order to elucidate the roles of enzyme and substrate ionizations, the effect of pH on the kinetic parameters of the demethylation reaction was determined throughout the pH range of 3.0 to 7.0, and the results were analyzed as described by Tipton and Dixon (14). Initial rate and inhibition studies of the CPO-catalyzed N-demethylation of DMA indicated that the reaction proceeds by a Ping-Pong kinetic mechanism (5). Thus, for the demethylation reaction, the I’,,,,,/ Km for each substrate (EtOOH or DMA) is the second-order rate constant for the binding and reaction of each substrate with the appropriate enzyme species (native CPO or compound I), and the pH dependence of the V,,,,,/K, will indicate kinetically important ionizations of the free substrate or enzyme species (14). Similarly, the pH dependence of the V,, will indicate kinetically important ionizations of the ES complex (14), since the I’,,,,, is the rate of the reaction at saturating concentrations of both substrates. For the demethylation reaction, the pH dependence of the V,,, will reflect ionizations on whichever ES complex (CPO-EtOOH or compound IDMA) is rate-limiting within the pH range. 8
8-
- 0.2
BY
-I
0.8 [Ethyl
Hydroperoxidel-‘(mM)-’
FIG. 1. Double-reciprocal plots of the initial rate of the demethylation of DMA at pH 4.5. The 3-ml reaction mixtures contained 0.5 M sodium potassium phosphate buffer, pH 4.5, and 0.29 yg CPO. (A) The concentration of DMA was varied as indicated in the presence of 5.36 mM EtOOH. (B) The concentration of EtOOH was varied in the presence of 5.26 mM DMA. The reactions were initiated by addition of the enzyme, and incubated for 10 min at 25’C. Formaldehyde formation was determined by the Nash assay as described under Experimental Procedures.
318
KEDDERIS
AND
The apparent kinetic parameters for the demethylation reaction as a function of pH are shown in Table I. The V,,, for the demethylation reaction exhibited a pH optimum of approximately 4.5. This is similar to the pH optima observed for the CPOcatalyzed evolution of oxygen from hydrogen peroxide (2) and for the N-oxidation of pchloroaniline (15). As shown in Table I, the Km for DMA decreased with increasing pH while the K, for EtOOH varied with pH in a manner paralleling V,,. The pH optimum of the V,,, for the demethylation reaction (pH 4.5) is different from the pH optimum of 6.0 previously reported (4). In the previous study (4), substrate concentrations which were saturating at pH 6.0 (0.53 mM DMA and 1.66 mM EtOOH) were used for the entire pH range, and the assumption was made that the initial velocities measured as a function of pH were proportional to the maximal velocities (14). However, it is apparent from the data in Table I that the decrease in the rate of the demethylation reaction with decreasing pH under the conditions used previously (4)
TABLE
I
EFFECT OF pH ON THE KINETIC PARAMETERS FOR CHLOROPEROXIDASE-CATALYZED DEMETH~ATION OF N,N-DIMETHYLANILINE’
K,,, DMA
PH 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0
@M) 23.27 13.57 5.36 3.05 0.809 0.382 0.088 0.061 0.034
f f f + It k * f +
K,,, EtOOH
(mM) 0.76 0.72 0.73 0.10 0.005 0.034 0.007 0.005 0.004
1.34 1.66 2.20 2.53 2.01 1.77 0.889 0.855 0.498
+ * f 5 I+ f * f k
0.01 0.03 0.10 0.09 0.23 0.05 0.022 0.021 0.063
V,, (nmol formaldehyde min-’ pg CPO-‘)
56.4 104.0 127.9 160.4 141.5 132.8 81.7 82.7 57.7
k f f + + k f + +-
6.6 13.6 3.3 5.6 9.0 8.1 12.8 5.4 6.0
“The reaction mixtures in sodium potassium phosphate buffer (0.5 M), pH as indicated, were initiated by addition of chloroperoxidase, incubated at 25°C for the times indicated under Results, terminated by addition of 60% trichloroacetic acid, and assayed for formaldehyde formation as described under Experimental Procedures. The chloroperoxidase concentrations were 0.895 ~3 at pH 3.0, 0.29 ~3 at pH 3.5 to 6.5, and 0.45 pg at pH 7.0. The data are presented as the mean values plus or minus standard errors from at least two determinations of K, and at least four determinations of V,,.
HOLLENBERG
was due to the use of subsaturating concentrations of DMA and EtOOH at pH values less than 6.0 rather than due to a ratecontrolling ionization. The data in Table I also indicate that CPO can efficiently demethylate both the protonated and neutral forms of DMA, suggesting that, under these conditions of ionic strength, hydrophobic binding of the DMA phenyl ring is more important in catalytic binding than ionic binding of the amine moiety. Comparison of the Vm,,/Km values for DMA and EtOOH (shown in Table II) indicates that the interaction of DMA with compound I is rate-limiting below pH 4.5, while compound I formation is rate-limiting above pH 4.5. Therefore, the pH dependence of the V,, for the demethylation reaction will reflect ionizations on the compound I-DMA complex at pH values below 4.5 and ionizations on the CPOEtOOH complex at pH values above 4.5. The V,,,,,/K, for DMA is approximately 700 times greater at pH 7.0 than at pH 3.0. In contrast, the V,,,/K, for EtOOH changes by less than a factor of three as the pH increases from 3.0 to 7.0. The kinetic constants in Table I were analyzed as described by Tipton and Dixon (14) in order to identify kinetically important ionizations of the free substrates and enzyme species. As shown in Fig. 2, the plot of the log of V,,,/K, versus pH was essentially independent of pH for EtOOH. Since the V,,,,,/K, for EtOOH represents the second-order rate constant for the reaction of EtOOH with CPO to form compound I, these results indicate that compound I formation was not affected by any ionizations in the pH range studied. The curvature in the plot at the extremes of pH is not great enough to indicate any ionizations, and is probably due to rate-controlling ionizations outside of the pH range studied. The V,,/K, for DMA represents the second-order rate constant for the reaction of DMA with CPO compound I. The plot of the log of the V,,,/K, for DMA (Fig. 3) increased with increasing pH in a linear fashion (slope = 1) through pH 6.0 and then leveled off. Extrapolation of the straight-line portions of the plot indicated
pH
STUDIES TABLE
ON
N-DEMETHYLATION
BY
319
CHLOROPEROXIDASE
II
EFFECT OF pH ON THE V/K VALUES FOR CHLOROPEROXIDASE-CATALYZED DEMETHYLATION OF N,N-DIMETHYLANILINE” V,,,,,/K,,, (nmol formaldehyde min-’ CPO-’ ITIM-‘)
fig
IO
PH
3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5
1360
7.0
1700
23.9
52.6 175 348
from
/ PH
FIG. 3. Dependence of the log of the V,,,,/K,,, for N,N-dimethylaniline upon pH. The VmI,../K,,, was calculated from the data in Table I. The standard errors in the V,,,,,/K,,, were determined by the propagation of error in the kinetic parameters.
92.0 96.7
928
a Calculated
05
42.1 62.7 58.1 63.4 70.4 75.0
2.42 7.66
116
the data
in Table
I.
a pK of approximately 6.8 on compound I or DMA. Since the pK of DMA is around 5.0 (lo-13), this must be an ionization of compound I. Thus, the ionization of DMA, although occurring in this pH range, is not important for catalysis since it does not show up in the kinetic plots. If the increase in the Km for DMA with decreasing pH (Table I) was due to its protonation to a form which could not be demethylated by the enzyme, then redefinition of the Km in terms of the concentration of the neutral species of DMA at a given pH should result
1,
I 30
1 40
I
1 50 PH
I
I 60
I
I 70
FIG. 2. Dependence of the log of the V,,/K, for ethyl hydroperoxide upon pH. The V,,/K,,, was calculated from the data in Table I. The standard errors in the V,‘,,/K,,, were determined by the propagation of error in the kinetic parameters.
in the pH independence of the Km. The Km can be redefined in terms of the concentration of the neutral species of DMA by the following modification of the Henderson-Hasselbach equation (16) Kk = K,/[l
+ antilog(pK
- pH)],
where Km is the apparent Michaelis constant at a given pH value, pK is the ionization constant of DMA (5.0), and K& is the Km in terms of the concentration of the neutral species of DMA. When this equation was used to redefine the apparent Km values for DMA in terms of the concentration of the neutral species of DMA, the redefined Km values were still pH dependent (not shown), further supporting the conclusion that the ionization of DMA is not catalytically important in the demethylation reaction. Extrapolation of linear tangents to the curved section of the plot of the log of V,,, versus pH (Fig. 4) at the lower limit of the pH range studied indicated a pK of approximately 3.1 on the compound I-DMA complex. The decrease in the log of V,,, at higher pH was not great enough to indicate a pK within the pH range studied, and is probably due to an ionization of the CPO-EtOOH complex with a pK greater than 7.0. This ionization is probably the
320
KEDDERIS
AND
244
2ot T---d 04I 30
1
I 40
1
1 50 PH
I
1 60
I
I 70
FIG. 4. Dependence of the log of the V,,,, for the demethylation reaction upon pH. The log of V,, was calculated from the data in Table I.
same group whose ionization in compound I was indicated in the plot of the log of Vm,,/Km for DMA versus pH (Fig. 3), its pK being shifted to a slightly higher value in the CPO-EtOOH complex. Similarly, the pK of approximately 3.1 indicated in the plot of the log of V,,,,, versus pH (Fig. 4) may be the same group responsible for the downward trend at low pH in the plot of the log of V,,,/K, for EtOOH versus pH (Fig. 2), its pK also being slightly higher in the compound I-DMA complex. This ionization may represent the transition between the acid and neutral forms of CPO, which has been reported to have a pK of approximately 3.5 (17). Hager and co-workers (18) have demonstrated that carbon monoxide binding by ferrous CPO is influenced by an ionizable group on the enzyme which exhibits a pK of approximately 5.5. The unprotonated form of the enzyme reacts with carbon monoxide much faster than the protonated form. Based on the pK and the AH“ (8 kcal mall’) for the ionization of the functional group affecting the binding of carbon monoxide, they suggest that this ionizable group is a histidine residue at the active site which is on the distal side close to the sixth axial ligand position of the heme iron atom. The studies described here on the CPO-catalyzed demethylation reaction do not provide any evidence for the involvement of a residue on the protein having a pK of 5.5. However, these studies involve the ferric as well as higher oxidation states
HOLLENBERG
of the heme iron, whereas in the carbon monoxide binding studies only the ferrous form could be investigated. Since variations in the electronic environment of amino acid residues in proteins may result in shifts in pK values of as much as 2-3 units (14), the ionizable group having a pK of approximately 6.8 which we observed in compound I may be the same imidazole histidine which has been shown by Hager and co-workers to play a role in the binding of carbon monoxide to ferrous CPO. However, the identities of the ionizable groups cannot be determined from the kinetic data presented here. The results of the pH kinetic analysis of the CPO-catalyzed demethylation reaction indicate that for optimal catalysis, a residue on the compound I-DMA complex with a pK of approximately 3.1 must be deprotonated while a residue on compound I with a pK of approximately 6.8 must be protonated. Marked substrate inhibition of the demethylation reaction by DMA was observed above pH 6.0 (data not shown), suggesting that deprotonation of the residue with a pK around 6.8 results in a higher affinity of DMA for an inhibitory site on the enzyme. The deprotonation of the group with a pK around 3.1 and the protonation of the group with a pK around 6.8 may be necessary to achieve the optimal conformation of the active site for catalysis. Over the pH range from 3.5 to 6.0, the two residues are mostly in their optimal ionization states for catalysis. At the extremes of the pH range, the active site may be in less optimal conformations, resulting in a slower rate of demethylation. ACKNOWLEDGMENTS We thank Professor Lowell P. Hager for supplying the purified chloroperoxidase and for his helpful suggestions and encouragement. We thank Ms. Nancy Starks for her excellent assistance in the preparation of this manuscript.
REFERENCES 1. MORRIS, D. R., AND HAGER, L. P. (1966). J. Biol. Chmz. 241, 1763-1’768. 2. THOMAS, J. A., MORRIS, D. R., AND HAGER, L. P. (1970) J. Biol. Chem 245, 3129-3134.
pH
STUDIES
ON
N-DEMETHYLATION
3. CORBEIT, M. D., BADEN, D. G., AND CHIPKO, B. R. (1979) B&n-g. Chm 8, 91-95. 4. KEDDF.RIS, G. L., KOOP, D. R., AND HOLLENBERG, P. F. (1980) J. Biol Chem. 255, 101’74-10182. 5. KEDDERIS, G. L., AND HOLLENBERG, P. F. (1983) J. BioL Chcm 258, 12413-12419. 6. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol Chem 193, 265-2’75. 7. SILBERT, L. S., AND SWERN, D. (1958) Ad Chem. 30,385-387. 8. NASH, T. (1953) Biochem J. 55,416-421. 9. EVERETT, A. J., AND MINKOFF, G. J. (1953) Trans. Faraday Sot. 49,410-414. 10. GUTBENZAHL, B., AND GRUNWALD, E. (1953) J. Amer. Chem. Sot 75, 559-565. 11. BACARELLA, A. L., GRUNWALD, E., MARSHALL, H. P., AND PURLEE, E. L. (1955) J. Org. Chem 20, 747-762.
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CHLOROPEROXIDASE
12. FICKLING, M. M., FISCHER, PACKER, J., AND VAUGHN, Chem Sot. 81.4226-4230. 13. CHO, A. K., AND MIWA, LXsp. 2, 477-483.
A., MANN, B. R., J. (1959) J. Amer.
G. T. (1973)
Drug
Met&
14. Tipton, K. F., and Dixon, H. B. F. (1979) in Methods in Enzymology (Purich, D. L., ed.), Vol. 63, pp. 183-234, Academic Press, New York. 15. CORBEIT, M. D., CHIPKO, A. 0. (1980) Biochem
B. R., AND BATCHELOR, J. 187, 893-903.
16. MCEWEN, C. M., JR., SASAKI, G., AND LENZ, W. R., JR. (1968) J. Biol Chew 243, 5217-5225. 17. THOMAS, (1970)
J. A., MORRIS, J. Bid Chem.
D. R., AND HAGER, 245, 3135-3142.
L. P.
18. CAMPBELL, B. A., JR., ARAISO, T., REINISCH, L., YUE, K. T., AND HAGER, L. P. (1982) Biochemistry 21, 4343-4349.