The temperature dependence of the components of the hepatic microsomal mixed-function oxidases

ARCHIVES

The

OF

BIOCHEMISTRY

AND

Temperature

BIOPHYSICS

Dependence Microsomal

JORDAN Clinical Department

227-234

160,

of the

AND

9, 1971;

of the

Hepatic

Oxidases MATTHEW

L. CARR’

Hospital, &linneapolis, Minnesota 66417,the Minneapolis, Minnesota 66466; and the Baltimore institute, Baltimore, Maryland 21211

Administration Pharmacology Unit, Veterans of Pharmacology, University of Minnesota, Cancer Research Center, National Cancer December

Components

Mixed-Function

L. HOLTZMAN

Received

(1972)

accepted

February

20, 1972

The activation energies of ethylmorphine N-demethylase (16.8 kcal/mole) and aniline hydroxylase (17.6 kcal/mole) did not greatly differ. The endogenous NADPH oxidase (16.8 kcal/mole) and the oxidase in the presence of ethylmorphine (15.7 kcal/mole) and aniline (14.7 kcal/mole) are only slightly different. On the other hand, t,he activation energy for NADPH-cytochrome c reductase is only 8.53 kcal/mole, suggesting that the reduction of the flavin in this enzyme does not significantly contribute to the rate of the overall reaction. The addition of ethylmorphine reduces the NADPH-cytochrome P-450 reductase activation energy from 12.5 kcal/mole to 10.8 kcal/mole. This energy is slightly increased by the presence of aniline (14.7 kcal/mole). Since NADPH-cytochrome P-450 reductase in the presence of ethylmorphine is presumed to be the rate-limiting step in the N-demethylase, the large difference between the energy of activation of this reaction and the energy of activation of ethylmorphine Cdemethylase would suggest the participation of two reactions of similar rate in the control of the overall rate of the oxygenase reaction.

The basic components of the enzyme complex involved in the hepatic, microsomal, mixed-function oxidases have been well elucidated over the past few years (1, 2). The reaction scheme involves a two-electron rcduction by NADPH of a flavoprotein, NADPH-cytochrome c reductase, which in turn reduces the cytochrome P-450-substrate complex. The reduced enzyme-substrate complex can then bind and reduce oxygen to give the hydroperoxide. The hydroperoxide can be thought of as disproportionating to give OH- and OH+, the latter being the so-called active oxygen which actually hydroxylates t,he substrate (3-5). At the present time, it is felt that the rate-limiting step in this series of reactions is the reduction of the cytochrome P-450 (6-8). This reduction can transfer only a single elect,ron to the iron of the heme. The ‘Present address: Department of Emory University School of Medicine, Georgia. Copyright

@ 1972 by Academic

Press,

ferroheme can then bind oxygen and tautomerize to give a ferriheme-superoxide complex (9-11). [Fez+

-

021 +

[Fe3+

-

02-I

In order to further reduce the oxygen to the hydroperoxide, the superoxide can either disproportionate (12) or receive a second electron from some other reduced species.Since this last step is irreversible, any reaction occurring after this during the steady-state process,as the attack of the substrate by the OH+, will have little or no effect on the rate. Since the NADPH-cytochrome P-450 reductase is the rate-limiting step in the mixedfunction oxidase activity, it could easily offer an optimal point for the control of “active oxygen” formation. And, indeed, Gigon et al. (13, 14) observed that the transport of the first electron to the cytochrome P-450, presumably for the formation of the ferrihemesuperoxide ion complex, is controlled by, or coupled to, the presenceof type I substrates.

Medicine, Atlanta,

Inc.

227

228

HOLTZMAN

They, as well as we and other workers, observed that these substrat’es could activate the reductase so that, the initial rate of rcduction of cytochrome P-450 could as much as double. Further, the difference between the rate in the presence and absence of the activator was, at 37°C for a one-elect’ron transfer, stoichiometric with the rate of hydroxylation (13-17). In confirmation of the importance of this tight coupling in the regulation of hydroxylation, we noted that in DzO there was a specific inhibition of the coupling, which was equal to the inhibition of the hydroxylation reaction (18). If, indeed, this coupled transport of electrons to reduce cytochrome P-450 is the rate-limiting process in the overall hydroxylase, then one would anticipate that the activation energy for hydroxylation should be comparable to that for the coupled reduction. We fully recognize that this assertion is hazardous in such a complex system, but when the various assays are run on the same preparation we should anticipate similar values. Yet, when we examined the temperature dependence of the various components of the hepatic mixed-function oxidases under conditions as close as is possible to those used in the overall reaction, we did not find good agreement. We interpret our results to indicate that the donation of the second electron significantly slows the rate of the reaction and probably comes from a second cytochrome P-450. METHODS In all experiments, 200- to 300-g fed, untreated, male S-D rats obtained from Charles River were used. The animals were killed by cervical fracture, the livers removed, chilled on ice, and homogenized in 3 ml of KC1 (150 mrvf-Tris (20 mM, pH 7.4) per g liver wet weight. The homogenate was centrifuged at 9000g for 15 min in either a Sorval RCP-B or International Equipment Corporation B-20 centrifuge. The 9000g supernatant fraction was centrifuged at 165,000g (av) in a Spinco Model L3-50 centrifuge for 38 min. The microsomal pellet was resuspended to the equivalent of 2 g of liver per ml of KCI-Tris. Protein was determined by the method of Sutherland et al. (19). All assays were performed in 3 ml of KC1 (150 mM)-Tris (20 mM, pH 7.4)-MgCl, (5 mivf) with tem-

AND

CARR

peratures of 1540%. When ethylmorphine and aniline hydrochloride (neutralized) were used, they were added as a 0.15 M solution to the buffer to give varying concentrations. In the assays of ethylmorphine N-demethylase and aniline hydrosylase, NADP (1. 2 pmoles), glucose-G-phosphate (30 pmoles) and glucose-6-phosphate dehydrogenase (2 units)2 were added. In all other enzyme assays NADPH (10 11, 100 mg/ml) was used. Microsomes were added to give a final concentration of 1 mg of protein per ml of incubate, except in the assay of NADPH-cytochrome c reductase and NADPH-cytochrome P-450 reductase where 25 pg and 3 mg of protein per ml, respectively, were used. Ethylmorphine X-demethylase and aniline hydroxylase were assayed in a shaking incubator under air, and the temperature maintained with a refrigerated circulating bath equipped with both a suction and an ejection pump. The times of incubation were 20 min at 15 and 2O”C, 15 min at 25”C, and 10 min at 30,35, and 40°C. The incubation with ethylmorphine was terminated by the addition of 1 ml of 5yG ZnSO,, 1.5 ml of BaOH (sat soln) and 0.5 ml NazBd07 . 10 Hz0 (sat soln). Aminophenol was extracted from 2.6 ml of aniline incubate with ethyl ether (25 ml) after adding solid NaCI. The formaldehyde was assyed in 4 M ammonium acetate with 0.4 ml of acetylacetone per 100 ml (20). Paminophenol was determined as the indophenol (6). The Km an d T’were determined by inspection. NADPH oxidase and NADPH-cytochrome c reductase activities were determined in an Aminco-Chance spectrophotometer (6). The initial rates were estimated by inspection. The NADPH oxidase rates are the average of duplicate incubations while the NADPH-cytochrome c reductases are the average of quadruplicate incubations. NADPH-cytochrome P-450 reductase activity was assayed with an Aminco anaerobic cuvette. The microsomal suspension either alone or with the ethylmorphine or aniline added was gassed for 5 min in a stream of CO gas deoxygenated with alkaline sodium hydrosulfite (8). The cuvette was capped after adding NADPH to the mixing plunger. The gas phase was flushed for 1 min and the cuvette placed in an Aminco-Chance spectrophotometer. The cuvette was allowed to equilibrate for 5 min (15, 20, 25, and 30°C) or 7 min (35 and 40°C). The initial rate was determined from the difference in absorbency between 450 and 490 nm. The rates are the average of duplicate incubations. 20ne unit at 25°C.

reduced

1 pmole

of NADP+

per minute

ACTIVATION

ENERGIES

OF HEPATIC

150 ' c

0

5 IO 15 20 25 30 35 40 45 50 55 Go Minutes

FIG.

la

MIXED-FUNCTION

OXIDASES

229

another type I substrate, aminopyrine. On the other hand, aniline, a type II substrate, did seem to show some differences in K,,, at, the lower temperatures (15.9 and 24.6”C). Due to the low activities at these temperatures it is unclear as to whether these differences are real or not. The Arrhenius plot of the V for the two activities are shown in Fig. 3. The activation energy (EJ for ethylmorphine N-demethylaseis 16.8 kcal/mole and aniline hydroxylase is 17.6 kcal/mole. Although a number of lines could be drawn through these points, those with the highest and lowest slopes differ from the mean Ea values by only 0.7 kcal/ mole, suggesting that these values have this level of precision. Repeated determinations have fallen within these bounds. If we examine the Arrhenius equation V = Ae-Wn/RT)

where V is the velocity at temperature T, EA the activation energy and R the gas constant, we find that it has a term (A) which varies as the v’/T and includes some term which is a function of the number of sites participating in the reaction (22). This term I I I , I I 0 5 10 15 20 25 30 35 40 45 50 55 60 has a value at 37°C of 6079 moles of HCHO formed/min/mg protein for ethylmorphine Minutes N-demethylase and 2285 moles of p-aminoFIG. lb phenol formed/min/mg protein for aniline FIG. 1. Time course of hepatic microsomal (a) hydroxylase. These crude values would apethylmorphine N-demethylase and (b) aniline hydroxylase from a male rat at various temperatures pear to have some significance in that they between 15 and 40°C. Preparations and assays are are not wildly different, contrary to our pregiven in the text. vious report (23) and would suggest that there are not grossly different concentrations RESULTS of the rate-limiting speciesfor the two reacAs can be seenin Fig. la and b, the rate of tions. The reason for the difference between both ethylmorphine N-demethylase and ani- our two studies is that in our previous study line hydroxylase is essentially linear for at the aniline hydrochloride solution was not fully neutralized and, therefore, the EA and least 10 min at all temperatures. Unfortunately, at the lower temperatures with the resultant A for aniline hydroxylase were lower concentrations of substrates, it is nec- obtained at pH 6.5 rather than 7.4. essary to use longer incubation times, even The activation energies of endogenous though the velocities are not precisely equal NADPH oxidase (16.8 kcal/mole), the oxito the initial velocity, in order to obtain dase in the presence of ethylmorphine (15.7 sufficiently high sensitivity in the assays. kcal/mole) and aniline (14.7 kcal/mole) are all comparable to the energy of the mixedDouble-reciprocal plots of ethylmorphine N-demethylase activity at varying temperafunction oxidases (Fig. 4). Since the entures indicate no major effect on the K, (Fig. dogenous, ethylmorphine-stimulated and 2a). This is not in agreement with the re- aniline-inhibited NADPH oxidase activation sults of Schenkman and Cinti (21) for energiesare fairly close,it would appear that

230

HOLTZMAN

AND

CARR

I/S (mM Eihylmorphind

FIG.

2a

L -30 l/S

[mM Aniline;-’

FIG.

FIG. 2. Double-reciprocal and (b) aniline hydroxylase Preparations and assays

2b plots of hepatic microsomal from a male rat at various are given in the text.

all three have a similar rate-limiting step. Further, since the rate-limiting step for NADPH-cytochromc c reductase is the reduction of the flavin (6, 24), it is clear that with an activation energy of 8.53 kcal/mole this reduction is not rate-limiting for the NADPH oxidase (Fig. 5). This difference between the energy of the NADPH-cytochrome c reductase and endogenous NADPH oxidase, would indicate that at best only a small fraction of the endogenous oxidase activity is due to direct oxidation of this flavoprotein by molecular oxygen. Therefore,

(a) ethylmorphine N-demethylase temperatures between 15 and 10°C.

either this flavoprotein must reduce oxygen through a cytochrome or other flavoproteins, with higher activation energies are involved in the NADPH oxidase activity. The relative, but not absolute insensitivity of the basal NADPH oxidase to CO would suggest that the reaction simult.aneously goes both by direct oxidation of flavoproteins and of cytochrome P-450 (6). The activation energy of the basal NADPH-cytochrome P-450 reductase activity (12.5 kcal/mole) is lower than the activation energy of either ethylmorphine

ACTIVATION

ENERGIES

OF-HEPATIC

MIXED-FUNCTION

OXIDASES

231

FIG. 3. Arrhenius plot of hepatic microsomal ethylmorphine N-demethylase (O--O) and aniline hydroxylase (O -----0) from a male rat. Values for the activity at each temperature were obtained from the estimated Vvalues observed in the plots shown in Figs. 2a) and b).

c cufference c An~llne InhIbited

I/T

(“K x IO ‘1

Fro. 4. Arrhenius plot of the endogenous (O--O), ethylmorphine-stimulated (A-----A), and aniline-inhibited (A-----A) NADPH oxidase of hepatic microsomes from male rats for temperatures between 15 and 40°C. The difference in activity between basal and ethylmorphine-stimulated oxidase activity is also shown (O--O). Preparations and assays are given in the text.

N-demethylase or aniline hydroxylase (Fig. 6). The addition of ethylmorphine reduces the energy to a value (10.8 kcal/mole) which is lessthan the energiesof either the endogenous reductase or ethylmorphine N-demethylase. Further, if one treats the difference between the endogenous and the ethylmorphine-stimulated reductase activities as if it were a separate enzymatic activity whose substrate is not reduced in the absence of ethylmorphine, we find that the activation

energy for it is only 6.18 kcal/mole. The value for the activation energy of the NADPH-cytochrome P-450 reductase in the presence of aniline (14.7 kcal/mole) is slightly higher than either the basal or the ethylmorphine-stimulated reductase. DISCUSSION

The observed difference in activation cnergy between ethylmorphine N-demethylase and NADPH-cytochrome P-450 reductase

232

HOLTZMAN

AND CARR

I/T kK x IO+]

FIG. 5. Arrhenius plot of hepatic microsomal NADPH-cytochrome c reductase from a male rat for temperatures between 15 and 40°C. Preparations and assays given in the text.

FIG. 6. Arrhenius plot of the endogenous (O--O), ethylmorphine-stimulated (A-----A) and aniline-inhibited (A-----A) NADPH-cytochrome P-450 reductase of hepatic microsomes from a male rat for temperatures between 15 and 40°C. The difference in activity between the basal and ethylmorphine-stimulated reductase is also shown (O--O). Preparations and assays given in the text.

would appear

with our that the rate-limiting process of the mixed-function oxidases is the difference between the activity of the NADPH-cytochrome P-450 reductase in the absence and presence of substrate (13-18). We fully recognize that this discrepancy may be an artifact of the system. In kinetic studies of microsomal enzymes, the tacit assumption is made that, although the components of the system are buried in a membrane, the observed activities can be analyzed by the use of equations derived for purified, soluble systems. It is possible that these differences may result from the interaccurrent

concept

to be at variance

tion of the various components under the slightly different experimental conditions for each of the assays. Yet these assays are so similar that this seemsunlikely. We feel that the activation energy for the reductase activities is genuinely far too low to be the sole rate-limiting step in the reaction. It is so low, in fact, that this activation processprobably represents only a conformaConal change in some fraction of the cytochrome P-450 protein, so that the altered cytochromes become more readily reducible. On the basisof these results, it would appear that the stoichiometry of this stimulation with the formation of formaldehyde is a

ACTIVATION

ENERGIES

NADPH’

OF HEPATIC

Reduced NADPH,--C&chrome

c-.

iNAurnGytochroma

CJ

MIXED-FUNCTION

OXIDASES

233

\-Fo?t~-45

Raductasa

NADPH ,

NADP

Oxidized .,.S”,# NYYY”tytochroma Raductosa

Reduced NADPHCytochroma Raductosa

FIG. 7. Schemefor the hepatic mixed-function oxidases.

fortuitous result associated with the temperature of the test solution. One interpretation of these results may be that t,he reduct,ion of the cytochrome P-450substrate complex is not the solerate-limit,ing factor in hydroxylations.3 On the basis of our present models of the microsomal mixed-function oxidases, there are four postulated reactions which, along with the reduction of the cytochrome P-450, occur before the formation of the hydroperoxide and may, therefore, affect the rate of the reaction. Two of these, the reduction of the flavin and the binding of the molecular oxygen to the ferroheme, are much too rapid to be important in determining the rate. The third, the activation of the reductase by the substrate, probably is not significant since the activation sites for this processare fully saturated at this concentration of substrate (15, 16). The fourth, the addition of the second reducing equivalent to the superoxide, therefore, must be making some contribution to the activation energy of the overall process. More recently, Estabrook et al. (25, 26) have presented spectrophotomet.ric evidence to support the existence of an oxygenated intermediate which may represent one of the tautomeric forms of the ferroheme oxygen complex. If such is the case, then the fact that peaks can be ob-

served in the Soret region may indicate that there is a significant accumulation of this intermediat’e and that it does not rapidly break down. The observations of Elison et al. (27), Mitoma et al. (28), and Tanabe et aE. (29) that the deuteration of aliphatic carbons to be hydroxylated gives a significant isotope effect might suggest that the rupture of the C-H bond is actually the rate-limiting step in the overall process, and not the formation of the “active oxygen.” If such were the case, one would not expect the activation energies to be the same. However, recent experiments in our laboratory would indicate that this isotope effect is only 25 % and, therefore, too small to support this hypothesis (Thompson, unpublished results). If the donation of the second electron does significantly affect the rate, then our data may give some clue as to the nature of the donating species. As has been previously discussed(6)) the secondreducing equivalent probably does not come directly from NADPH, or through a sulfhydro group, nonheme iron protein, or a flavoprotein. The two most likely candidates would, therefore, be either cytochrome bg (30) or a second cytochrome P-450 bound to oxygen. In order to evaluate the possible importance of cytochrome bbin this reaction, we attempted to measure the activation energy of the re3It shouldbe noted that Schenkmanand Cinti duction of this cytochrome. In the presence (21) did not report any suchdiscrepancybetween of air and using either NADPH or NADH, the reductaseanddemethylaseenergiesfor aminopyrine. The reasonfor this differencemay againbe the reduction at 25°C was complete in intact microsomesin less than a half second. With due to the useof a different substrate.

234

HOLTZMAN

such a rapid reaction, the reduction of this species could not contribute to the activation energy. It would, therefore, seem most likely that the second electron is derived from a second reduced cytochrome P-450 (Fig. 7).4 The rate of the overall reaction is, therefore, going to be altered by the rate of reduction of this second cytochrome. REFERENCES 1. GILLETTE, J. R. (1966) Advan. Pharmacol. 4, 245. 2. MANNERING, G. J. (1971) Fundamentals of Drug Metabolism (LaDu, B., Mandel, H. G., and Way, E. L., eds.) Williams & Wilkins, Baltimore. 3. HAYAISHI, O., AND NOZAKI, M. (1969) Science 164,389. 4. ULLRICH, V., .4ND STAUDINGER, H. (1969) Microsomes and Drug Oxidations (Gillette, J. R., Conney, A. H., Cosmides, G. J., Eastabrook, R. W., Fouts, J. R., and Mannering, G. J., eds.) Academic Press, New York. 5. GUROFF, G., DALY, J. W., JQRINA, D. M., RENSON, J., WITKOP, B., .~ND UDENFRIGND, S. (1967) Science 167, 1524. 6. HOLTZMAN, J. L. (1970) Biochemistry 9, 995. 7. OMUR.~, T., SANDERS, E., ESTABROOK, R. W., COOPER, D. Y., AND ROSENTHAL, 0. (1966) Arch. Biochem. Biophys. 117, 660. 8. HOLTZMAN, J. L., GRIM, T. E., GIGON, P. L., AND GILLETTE, J. R. (1968). Biochem. J. 110,407. 9. GUNS.~LUS, I. C., CONRAD, H. E., :\ND TRUDGILL, P. W. (1965) in Oxidases and Related Redox Systems (King, T. S., Mason, H. S., and Morrison, H., eds.) Wiley, New York. 10. ORME-JOHNSON, W. H., AND BEINERT, H. (1969) Biochem. Biophys. Res. Commun. 36, 905. 4Although this scheme may appear to call for two NADPH per hydroxylation, such is not actually the case. It is well known that the reduced flavin can transfer one electron to the cytochrome P-450 to become a free radical semiquinone. This can then transfer the second electron to a second cytochrome P-450 to give the proper stoichiometry.

AND

CARR

11. MISSEY, V., STRICKL.\ND, S., M.~YHF,M, S. G., HOWELL, L. G., ENGEL, P. C., MATTHEWS, It. G., SCHUhlSN, M., hND SULLIVAN, P. A. (1969) B&kern. Biopkys. Aes. Commun. 36, 891. 12. MO~:LLER, T. (1952) Inorganic Chemistry, Wiley, New York. 13. GIGON, P. L., GR.~M, T. E., J~ND GILLETTE, J. R. (1968) Biochem. Biophys. Res. Commun. 31, 558. 14. GIGON, P. L., GRAM, T. E., AND GILLETTE, J. R. (1969) Mol. Pkarmacol. 6, 109. 15. HOLTZMIN, J. L., AND RUMACK, B. H. (1971) Life Sci. 10, 669. 16. HOLTZMAN, J. L., .~ND RUMACK, B. H. (1971) Chem. Biol. Interactions 3, 279. 17. ULLRICH, V. (1969) 4th Int. Congr. Pharmacol. Basel. p. 14. 18. HOLTZMAN, J. L., AND CARR, M. L. (1970) Lije Sci. 9, 1033. 19. SUTHERL.~ND, E. W., CORI, C. F., HAYNES, R., AND OLSEN, N. S. (1949) J. Biol. Chem. 130, 825. 20. NASH, T. (1953) Biochem. J. 66,416. 21. SCHENKMAN, J. B., AND CINTI, D. L. (1970) Biockem. Pharmacol. 19, 2396. 22. MOORE, W. J. (1952) Physical Chemistry, p. 531, Prentice-Hall, New York. 23. HOLTZMAN, J. L., AND C~RR, M. L. (1970) Pharmacologist 12, 336. 24. K~MIN, H., MASTERS, B. S. S., GIBSON, Q. H., AND WILLIAMS, C. H. (1965) Fed. Proc. 24, 1164. 25. ESTIBROOK, R. W., BARON, J., END HILDEBRANDT, A. (1971) Fed. PrOc. 30.1729. 26. EST..\BROOK, R. W., HILDEBRANDT, A. G., B.IRON, J., NETTER, K. J., .~ND LEIBIXAN, K. (1971) Biockem. Biophys. Res. Commun. 42, 132. 27. ELISON, C., ELLIOTT, H. W., LOOK, M., AND R.APP.~PORT, H. (1963) J. Med. Ckem. 6, 237. 28. MI~POM:\, C., YASUDA, D. M., T.IGG, J., AND T~N~BE, M. (1967) Biockim. Biophys. Acta 136, 566. 29. TANABE, M., YASUDA, D., LEV.ILLEY, S., AND MITOM.~, C. (1969) Life Sci. 8.1123. 30. ESTARROOK, R. W., .SND COHEN, B. (1969) in Microsomes and Drug Oxidation (Gillette, J. R., Conney, A. H., Cosmides, G. J., Estabrook, R. W., Fouts, J. R., and Mannering, G. J., eds.), Academic Press, New York.