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Microsomal oxidase IV: Properties of a mixed-function amine oxidase isolated from pig liver microsomes
ARCHIVES
OF
BIOCHEMISTRY
Microsomal
AND
Oxidase Oxidase DANIEL
Clayton
Foundation
116-125
BIOPHYSICS
160,
IV:
Properties
Isolated M. ZIEGLER2
(1972)
of a Mixed-Function
from
Pig
Liver
CAROLYN
AND
Microsomes’ H. MITCHELL3
Biochemical Institute and the Department of of Texas at Austin, Austin, Texas Received
December
22, 1971;
accepted
Amine
February
Chemistry,
The
University
3, 1972
A flavoprotein-oxidase isolated from pig hepatic microsomes catalyzes the NADPHand oxygen-dependent N-oxidation of a variety of lipid-soluble set- and tert-amines. The set-amines are oxidized to the hydroxylamines; and the tert-amines, to the amine oxides. With one or two exceptions the isolated oxidase does not catalyze the oxidation of primary amines. However, primary alkylamines added to the reaction vessel increase the set- and tert-amine N-oxidase activity of both the isolated enzyme and microsomes. The isolated oxidase contains 14 nmoles FAD and about 0.02 mg lipid/mg protein. The oxidase appears to be free from cytochromes but is contaminated with very small, but detectable, amounts of a hemoprotein that corresponds to catalase, both spectrally and in enzymic properties. The isolated enzyme is free from NADPH-cyt. c reductase, diaphorase, and monoamine oxidase activities.
An earlier report (1) demonstrated that hepatic tissues from vertebrates contained an oxidase that catalyzed the oxidation of of DMA4 to the amine oxide. While this activity could be detected in all vertebrates tested, the DMA N-oxidase activity was consistently highest in hepatic tissue from pigs. The speciesand organ distribution of DMA N-oxidase activity resembled the distribution of the trimethylamine N-oxidase described by Baker et al. (2). Both DMA N-oxidase and the trimethylamine Noxidase require NADPH for activity, and both activities are concentrated in the microsomal fraction of liver homogenates. ‘This work was supported in part by U. S. Public Health Service Grant GM 12360. ZWork carried out during tenure of a U. S. Public Health Service Career Development Award (l-K3GM-25,990). 3California Institute of Technology, Pasadena, CA. ‘Abbreviations used are: DMA, N,N-dimethylaniline ; DMOA, N, N-dimethyl-n-octylamine; SKF-525A, Diethylaminoethanol ester of diphenylpropylacetate. 116 Copyright
@ 1972 by Academic
Press,
Inc.
Baker and Chaykin (3) have also shown that the oxygen atom incorporated into trimethylamine is derived from molecular oxygen. The enzyme catalyzing the Noxidation of DMA appeared to be of the type classified as mixed-function oxidases (4). Unlike other widely studied hepatic microsomal mixed-function oxidases (5)) the DMA N-oxidase was reported to be insensitive to both carbon monoxide and SKF-525A(6). This and other evidence suggested that cyt. P-450 was not an obligatory component of the microsomal DMA Noxidase system. Both in vitro and in vivo N-oxidation of other tertiary amines has been demonstrated in many vertebrates including man (1, 2,7-g). While N-oxides are generally considered minor metabolites of amine metabolism, the rapid N-oxidation of certain tertiary amine drugs has been reperted (cf. review by Bickel, 10); and Beckett and Hewick (11) have shown in vitro that amine oxides are the major oxidation products of the phenothiazine drugs.
MICROSOMAL
The N-hydroxylation of secondary amines has also been observed in vitro, and the extensive studies of Kiese and his associates (12, 13) have shown that the organ and subcellular distribution and the cofactor requirements of the secondary amine Nhydroxylase are virtually identical with those of the tertiary amine N-oxidase. Their studies support the interpretation that the same enzyme catalyzes the N-oxidation of both secondary and tertiary amines. This report describes the isolation from pig liver microsomes of a flavoproteinoxidase that catalyzes the NADPH- and oxygen-dependent N-oxidation of secondary and tertiary amines. Some of the properties of partially purified preparations of this oxidase were described in an earlier report (14). EXPERIMENTAL
aminesubstrates,obtainedfrom commercialsources,were purified by gas-liquid Materials.
The
chromatography and then stored at -15°C. As needed, aliquots of the pure amines were dissolved in equivalent amounts of 0.1 M HCl. Solutions of the aniline derivatives were prepared daily. Aqueous solutions of the other amines are more stable and were stored at - 15°C for several weeks without undergoing any apparent change. Ultrapure sucrose and ammonium sulfate were obtained from Mann Research Lab; Alphacel and protamine sulfate, from Nutritional Biochem. Corp. ; NADP+, NADPH, Sephadex G-100-120, DEAE-cellulose, Triton-X-45, Triton X-100, and Triton-X-102, from Sigma Chemical Company. In addition, samples of all the detergents used in this study were initally supplied by R,ohm, Haas and Company, Philadelphia. Ten-per cent solutions of each of the Triton detergents were prepared by dissolving 100 g of the detergent in water to a final volume of 1 liter. The solutions were stored at 0-5°C. Even though a mixture of Triton-X-102 and Triton-X-45 was used to extract the oxidase, these two detergents had to be stored as separate solutions. Equal volumes of the two detergent solutions were mixed just prior to use. The saturated “acid’‘-ammonium sulfate solution was prepared by dropwise addition of concentrated sulfuric acid to a solution saturated at room temperature with ammonium sulfate. Sufficient sulfuric acid was added to bring the pH of the solution to 4.8 f 0.1 at 25’C after it was diluted lo-fold with water. The acid-ammonium sulfate solution was stored at O-5’% for at least 1 day before use.
OXIDASE
IV
117
The pretreatment and conditions for storing DEAE-cellulose, Sephadex, and cellulose were as follows: The DEAE-cellulose was suspended in about 10 vol of 0.5 M HCl and then washedwith water until the effluent was free from chloride ions. The washed ion-exchange cellulose was resuspended in about 10 vol 0.03 M glycine and stored at 0-5°C. Fifteen grams of Sephadex G-100-120 suspended in 2 liters of water were heated in a steam cabinet for 8 hr. Before cooling, HCl was added to the hydrated Sephadex to a final concentration of 0.1 M; and the suspension was heated to boiling for 5 min. The mixture was rapidly cooled on ice, and the Sephadex was washed with cold water until free from chloride ions. The gel, suspended in 5 vol 0.05 M glycine, can be stored for several months at 0-5°C. The cellulose (Alphacel) was treated three to four times (alternately) with 0.02 M HCl and 0.02 M KOH to remove extraneous material. The cellulose was collected by filtration, washed with a large excess of water to remove the last traces of KOH (pH of the filtrate about 6), resuspended in water, and stored in the refrigerator until needed. Pig liver microsomes were isolated by the method described earlier (6). The liver tissue was obtained almost exclusively from old sows since, as shown with rats (15), the hepatic N-oxidase activity is higher per milligram of protein in tissue from older animals. Methods. Unless otherwise indicated, enzyme assays were carried out in open lo-ml Erlenmeyer flasks as described earlier (16). The reaction medium routinely contained 0.5 mm NADP+; 5.0 rnM magnesium chloride; 5 mM isocitrate; isocitrate dehydrogenase (sufficient to reduce 1 pmole NADP+/min/ml); and 0.1 M alanine-0.02 M pyrophosphate buffer, pH 8.4, at 38°C. After a 5-min temperature equilibration the reaction was started by the addition of enzyme. The amine oxidation products were measured as described earlier (1517). Protein and acid-extractable flavin were measured by standard methods (cf. Ref. 15). Lipids extracted by the Folch procedure from the purified oxidase were assayed for neutral lipids by the method of Brown and Johnston (18). The phospholipids were separated by tic, and the phosphate concentration of the individual phospholipids was measured by the method of Parker and Peterson (19). Isolation of the mixed-function amine N-oxidase. All steps in the procedure were carried out at O-5’ C. Microsomes stored in 0.25 M sucrose at - 15’C for no longer than 1 month were thawed, diluted to approximately 20 mg protein/ml, and adjusted to pH 5.8 by dropwise addition of 1 M acetic acid. The suspension was immediately centrifuged at 30,000 rpm for 10 min. The residue was collected and re-
118
ZIEGLER
PURIFIC.~TION
OF THE MIXED-FUNCTION
AND
MITCHELL
TABLE AMINE
I OXIDASE
Volume (ml)
Fraction
FROM Protein (w)
-_ ..Microsomes 3550%
Ammonium
sulfate
Acid-ammonium
sulfate
After
DEANE-cellulose
After After
Sephadex G-100-129 electrophoresis
(1 The assays was 2.5 mM; the b The values tions that have
chromatography chromatography
400
9600
120
830
16
150
8
118
2 3
75 26
PIG
LIVER
nmoles DMA oxidized min/mg
protein
23.0 (16-38)” 160 (99-210) . 520 (36&600) (47E20)
Percent of original activity 100 60 35 33
890 1,049
were carried out at 38°C as described in Methods. The initial reaction time was 3 min. listed in parentheses are the lowest and highest specific activities been successfully carried through the succeeding steps.
suspended in water to approximately 20 mg protein/ml and then was adjusted to pH 8.0 with dilute KOH. Four milliliters of 2.5 M guanidine hydrochloride were added for each 100 ml of the suspension, and the mixture wasthoroughly homogenized in a glass homogenizer fitted with a Teflon pestle. The aggregated particles were sedimented by centrifugation at 30,000 rpm for 15 min. The residue was resuspended in water and homogenized. The volume was adjusted to give a protein concentration of 29 f 1.0 mg/ml. The oxidase was extracted from the particles by adding, for each 100 ml of the suspension, 9 ml of the Triton X-45Triton X-102 solution (see Materials section). After stirring for 1 hr, 100 mg protamine sulfate were added for each 100 ml of the suspension. The correct amount of protamine sulfate was routinely dissolved in a minimum volume of water at 38°C and slowly added to the microsomal suspension with rapid stirring. The detergent-insoluble material was removed by centrifugation at 30,900 rpm in a Spinco Model L centrifuge for 1 hr. The clear, yellow-brown supernatant fraction was collected and immediately fractionated with ammonium sulfate. The fraction that precipitated upon the addition of 19.4 g ammonium sulfate for each 100 ml of solution was removed by centrifugation. This fraction collected as an oily pehicle at the top of t.he centrifuge tube and was removed and discarded. The soluble fraction was, if necessary, carefully filtered through glass wool to remove any remaining insoluble material; and an additional 8.7 g ammonium sulfate were added for each 100 ml of solution to precipitate the oxidase. The precipitate was collected by centrifugation and re-
MICROSOMEP
30 12
concentration of different
of DMA prepara-
dissolved in water to a final protein concentration of not more than 8 mg/ml. This fraction is referred to in Table I as the 35-5070 ammonium sulfate fraction. One-tenth volume of 1 M sodium acetate, pH 4.8, was added to the 35-5Oyc ammonium sulfate fraction; and the preparation was refractionated by adding the saturated acid-ammonium sulfate solution (see Materials section). Fractions precipitating at O-35,35-40,40-45, 4550% saturation were collected by centrifugation and redissolved in 0.05 M glycine, pH 6.2-6.6. The 0-35yc saturation fraction usually contained little or no amine oxidase activity and was discarded. The enzyme was usually most concentrated in the 4045% ammonium sulfate fraction while the other two fractions usually contained significant amounts of the oxidase. The latter two fractions (3540$& and 4550%) were combined, and sufficient saturated acid-ammonium sulfate was added to produce a faint turbidity. After stirring for 30 min, the precipitate was removed by centrifugation and discarded. The oxidase in the supernatant fraction was precipitated with ammonium sulfate (about 450/o saturation), collected by centrifugation, and combined with the initial 40-45’% saturation fraction. The combined fractions are referred to as the acid-ammonium sulfate fraction in Table I. Triton-X-100 (0.1 mg/ml protein) was added to the acid-ammonium sulfate fraction before it was carried through the final steps. After adding the detergent, the preparation was dialyzed with internal and external stirring against 20 vol water for 90 min. The water was changed every halfhour. The dialyzed preparation, in 10 ml or less,
MICROSOMAL was passed through a column (20 X 2.5 cm) of DEAE-cellulose that had been equilibrated with several column volumes of a solution containing 0.03 M glycine and 0.03 M histidine, pH 6.3. The oxidase, an intense yellow band, was washed through the column with 0.03 M histidine buffer, pH 6.3. The eluate containing the oxidase was reduced in volume to about 2 ml by pressure dialysis in an Amicon Diaflo Cell (Model 52) fitted with an XM-100 membrane. The concentrated fraction was immediately placed on a column (20 X 2.5 cm) of Sephadex G-100-120 equilibrated with 0.03 M glytine. Four-milliliter fractions were collected at a flow rate not greater than 0.5 ml/min. The fractions containing the oxidase were combined and reduced in volume to 1.5-2 ml by pressure dialysis. The final step in the isolation of the oxidase was carried out by zone electrophoresis in a unit similar to that described by Hauschild-Rogat and Smith (20) for preparative disc electrophoresis, except that the polyacrylamide gel was replaced with cellulose. The buffers used were the same as those specified by Davis (21) for polyacrylamide gel-disc electrophoresis. A maximum of 2 ml of each Sephadex G-100-120 fraction, containing Tris buffer, pH 6.7, was carried through the electrophoresis step at one time. In 5 hr at 350 V and 10 mA, the oxidase migrates 4-6 cm as a narrow, intensely yellow band. When the enzyme was about 2-3 mm from the bottom of the cellulose column, the electrodes were removed; and the oxidase was eluted with the electrode buffer at a flow rate not greater than 0.1 ml/min. The yellow fraction, in a volume of 2-3 ml, was collected and concentrated almost to dryness by pressure dialysis. The enzyme was taken up in 3-5 ml of 0.05 M histidine, pH 6.6-7.0, and stored at 0-3°C. Stability of the amine oxidase during isolation. The oxidase is quite unstable immediately after it is extracted from the particles, and all of the steps through the 35-500/, ammonium sulfate fraction must be carried out as rapidly as possible. The 35-50y0 ammonium sulfate fraction, if diluted to a protein concentration between 5-8 mg/ml, can be stored overnight at 0-5°C with very little loss of activity. If this fraction is stored at a higher protein concentration or for longer periods of time, progressive inactivation of the enzyme is frequently observed. In contrast, the oxidase in the acid-ammonium sulfate fraction is quite stable and can be stored at -15°C for several months with little or no loss of activity.After Sephadex chromatography the oxidase remains fully active when stored at 0-3°C for up to 2 weeks, but 20-40~0 of the activity is lost every time the preparation is frozen and thawed. Inactivation of the purified oxidase by freezing and thawing can be retarded, but not completely prevented, by the addition of 0.1 mg
OXIDASE
119
IV
Triton-X-lOO/mg enzyme. The most stable form of the oxidase is found in the acid-ammonium sulfate fraction, and the preparations are routinely stored at this stage. As needed, aliquots of this fraction are carried through the final steps in the isolation. RESULTS
The concentration of the oxidase varies widely in different preparations of pig liver microsomes (6). As a result the yield and concentration of the oxidase obtained during isolation were also extremely variable. If the initial specific activity of the microsomes is less than 15 nmoles DMA N-oxidized/ min/mg protein, the yield, st.ability, and purity of the oxidase are very erratic. Preparations of the oxidase at the highest level of purification are consistently obtained from microsomal fractions with an initial DMA N-oxidase specific activity of 18 or greater, and the concentration of the oxidase/mg protein obtained after the different steps is as indicated in Table I. The concentration of the enzyme after passage through DEAE-cellulose is not great and is somewhat variable, but this step is necessary to remove the last traces of cyt. bg. Any cyt. bgnot removed at this point will concentrate with the oxidase during chromatography on Sephadex. Both the pH and ionic strength of the amino acid buffer during DEAE-cellulose chromatography are critical. At a more neutral pH or lower ionic strength, the oxidase is irreversibly adsorbed on the ion-exchange cellulose. Irreversible binding of the enzyme on Sephadex G-100-120 does not occur; and at the low ionic strength used, virtually all of the purification of the oxidase may be attributed to the ion-exchange properties (enhanced by heating in dilute acid; cf. Methods section) of the Sephadex. In the presenceof 0.1 M KCl, very little purification of the oxidase is obtained by chromatography on Sephadex G-100-120. Composition of the oxidase. The purified oxidase contains 14 + 0.2 nmoles of flavin/ mg protein. All of the flavin can be ext’racted with acid and appears to be identical with FAD both spectrally and in its ability to activate the apo-D-amino acid oxidase. The amine oxidase contains about 2% lipid by weight. Analysis of the lipids extracted by
120
ZIEGLER
AND MITCHELL
the E’olch procedure from 23 mg of the purified oxidase showed the presence of the following in micrograms per milligram protein: phosphatidyl choline, 14.7; lysophosphatidyl choline, 2.7; free fatty acids, 3.9; unidentified phospholipids, 0.9; and small but detectable amounts of cholesterol. The purified oxidase appears to be entirely free from the detergents used to extract the enzyme from the particles. The characteristic uv absorption bands of the Tritons cannot be detected in dried acetone-extract’s of the oxidase redissolved in methanol or wat,er. Tritons added to the purified enzyme can be readily extracted and identified by this procedure. The oxidase does not contain detectable amounts of iron, copper, or cytochromes. The purified oxidase is also free from detectable NADH or NADPH diaphorase, cyt. c reductase, and monoamine oxidase activities. The purified preparation contains small but detectable amounts of catalase activity. Spectra. The spectrum of the oxidized flavoprotein (Fig. 1) is similar to that of other flavoproteins. The 260 nm/460 nm absorption ratio is greater than that of most flavoproteins. The absorption of the oxidized enzyme at 410 nm is also greater than expected for a pure flavoprotein. The increased absorption in this region of the spectrum may be due to the presence of small amounts of hemoprotein contaminant, probably catalase. Anaerobically, both the 380 and 450 absorption bands of the oxidase are completely reduced by NADPH; but in the presence of oxygen, NADPH produces no perceptible change in the spectrum of the oxidized enzyme. NSDPH oxidase activity. The isolated oxidase catalyzes the oxidation by oxygen of 90-110 nmoles of NADPH/min/mg protein at 38°C. A number of observations suggest that this activity is an inherent property of the oxidase and is not due to a contaminating NADPH-dependent enzyme. The pH optimum (8.4) for the NADPHoxidase activity is identical with the pH opt’imum for the N-oxidase activity of the enzyme. The ratio of the DMA-N-oxidase/ NADPH-oxidase activities remains relatively constant (10/l) during the last four
320 WAVELENGTH
400
460
560
my
FIG. 1. Direct spectraof the purified amineoxidase. The concentration of the enzyme is 0.8 mg protein/ml in 0.1 M glycine-O.02 M pyrophosphate, pH 8.4. -, oxidized enzyme; * * * * *, enzyme anaerobically reduced by 0.1 mM NADPH; and -----, enzyme reduced with NazS204. The spectrum of the NADPH-reduced enzyme was recorded after deaeration of the sample with nitrogen followed by addition of the isocitrate dehydrogenase-NADPH generating system. The same concentrations of NADPH and the generating system were added to the reference cuvette to minimize interference by the 340-nm band of NADPH with the spectrum of the reduced enzyme.
steps in the isolation of the oxidase. Attempts to differentiate between the two activities by pretreating the oxidase with heat detergents, mecurials, and proteolytic enzymes have been unsuccessful. Both activities are destroyed at the same rate. In the absence of an amine substrate the molar ratio of NADPH oxidized/Oz consumed, measured polarographically, is 2.0 (Poulsen, unpublished experiments this laboratory). Since the preparation also contains small quantities of catalase, these data suggest that hydrogen peroxide is produced during the reoxidation of the flavoprotein by oxygen. In the presence of saturating levels of added catalase, the purified oxidase will catalyze the peroxidation of methanol at about the same rate as that for NADPH oxidation, indicating that hydrogen peroxide is produced during the oxidation of NADPH.
MICROSOMAL
N-OXIDATION
TABLE OF AMINES PURIFIED AMINE
Substrate
II C.~T.~LYZED 0x1~~~~
nmoles oxidation product/min/mg protein N-Oxide
Tertiary amines Trimethylamine N, N-dimethylbutylamine N,N-dimethylhexylamine N, N-dimethylcyclohexylamine N,N-dimethyloctylamine N, N -dimethylnonylamine N,N-dimethyldodexylamineb N, N -dimethylaniline N, N -diethylaniline N-methylmorpholine N-ethylmorpholine Secondary amines N-methylaniline N-ethylaniline N-methyloctylamine
BY THE
Hydroxylamine
260 546 552 516
-
1650 1760 360
-
910 712 378 284
-
-
507 380 1020
a The fraction obtained after Sephadex G-lOO120 chromatography. Except for N-ethylmorpholine, the rates listed for all amine substrates were calculated from the rate of oxidation-product formation at 38°C in 2 min with 2.5 mnn amine substrate and 0.1-0.3 mg enzyme/ml. The rate for N-ethylmorpholine is based on the substratedependent oxidation of NADPH. b Concentration of this substrate was 0.5 mu.
Amine oxidation products. In addition to DMA, the purified amine oxidase catalyzes the N-oxidation of a number of other secondary and tertiary amines (Table II). The tertiary amines are oxidized to the corresponding amine oxides; and the secondary amines, to hydroxylamines. As shown earlier (22) with hepatic microsomes, the N, N-dimethyl-n-alkylamines with S-12 carbons in the side chain are N-oxidized at the fastest rates. The size, but not the arrangement, of the alkyl side cha,in appears to determine the rate at which the oxidase catalyzes N-oxidation of the substrate. For example, the N, N-dimethyl derivatives of n-hexyland cyclohexyl-amines are Noxidized at about the same rate. Amines
OXIDASE
IV
121
with N-ethyl groups are oxidized at slower rates than the corresponding N-methyl compounds (cf. derivatives of morpholine and aniline in Table 11). The secondary N-methyl amines are oxidized at about N the rate of the corresponding N, N-dimethyl tertiary amines. While only a limited number of secondary amines have been extensively studied, the effect of the side chain on the oxidation rate appears to be similar to that observed with the tertiary amines. Hydroxylamines appear to be the only oxidation products obtained from secondary amines. An earlier report indicated that a partially purified preparation of the amine oxidase catalyzed the oxidative N-demethylation of secondary amines (14). This interpretation was based on t’he observations that substantial amounts of formaldehyde can be detected in deproteinized aliquots of the reaction media containing the secondary amine substrates and the oxidase and t.hat the formation of formaldehyde was dependent upon substrate concentration and incubation time. However, it would appear that the formaldehyde is an aciddegradation product of the enzymically formed Nmethylalkylhydroxylamines. If aliquots of the reaction mixture are extracted with ether to remove hydroxylamines before deproteinizing with trichloroacetic acid, formaldehyde cannot, be detected in the aqueous phase. Formaldehyde added to the reaction vessels, with or without hydroxylamine, can be qualitatively recovered in the aqueous phase after repeated extraction with ether. The purified oxidase does not catalyze the oxidation of most primary amines or of secondary or tertiary amines with polar groups near the nitrogen. Oxidation of the N-methyl derivatives of acetanilide, formanilide, and nicotinamide could not. be detected. Stoichiometry. As indicated by the experiments summarized in Fig. 2, 1 mole NADPH is required for each mole of substrate Noxidized. The ratio of the rate of DMOAdependent NADPH oxidation to the rate of DMOA N-oxide formation is 1: 1. However, the rate of DMA N-oxide formation is somewhat greater than the rate of DMAdependent NADPH oxidation and is equal
122
ZIEGLER
AND MITCHELL
I 0 0
I
2 TlME
3
4
I 2
I 4
t 6
I
5 I/DMOAlmM)-'
(mill)
FIG. 2. Rates of NADPH oxidation and amine oxide formation. The reactions were carried out at 38°C in 3.0~ml cuvettes in a Zeiss PM-&II spectrophotometer. NADPH concentrations were measured at 340 nm, and aliquots of the reaction mixtures were withdrawn and analyzed for amine oxides at the times indicated. A-A, rate of DMOA-N-oxide formation. O-O, rate of DMOAdependent NADPH oxidation. A-A, rate of DMA-N-oxide formation. a-0, rate of DMAdependent NADPH oxidation. The initial concentrations of NADPH, DMOA, and DMA were 0.2, 3.0, and 1.5 mM, respectively.
to, or slightly less than, the total rate of NADPH oxidation. Thus, DMA, in contrast to DMOA, appears to block the endogenous NADPH oxidase activity of the isolated oxidase. E$ect of primary amines on activity. Double reciprocal plots of DMOA concentration versus reaction rate are shown in Fig. 3. The points do not fall on a straight line, and a pronounced concave curve is obtained over the range of DMOA concentrations tested. High concentrations of this substrate apparently activate the oxidase, and this activation is quite evident at DMOA concentrations above 1 mu. However, amines that are not substrates for the oxidase can also activate the enzyme. As shown in Fig. 3, the enzymic rate at low DMOA concentrations is almost doubled in the presence of 5 mu n-octylamine. At the highest concentration of DMOA tested, the addition of octylamine did not increase the oxidation rate of DMOA; and both curves in Fig. 3 appear to intersect the y axis at about the same point. In cont’rast to the alkylamines, the aniline
FIG. 3. l/DMOA vs. l/v, where v is the rate of DMOA-N-oxide formation/min/mg protein at pH 8.4 and 38°C. O-0, DMOA. wm, DMOA in the presence of 5.0 lllM n-octylamine.
‘,“I 1.0
-
z 0.5-
-40
-30
-20
-10
0
10
20
-I
l/DMA
(mM1
FIQ. 4. l/DMA vs. l/v, where v is the rate of DMA-N-oxide formation/min/mg protein at pH 8.4 and 38°C. +-@, DMA. H, DMA in the presence of 1.0 rnM n-octylamine.
derivatives do not activate the oxidase. As shown in Fig. 4, double reciprocal plots of DMA concentration versus rate yield straight lines. The addition of 1.0 mu n-octylamine doubles the rate of DMA N-oxidation over the whole range of DMA concentrations tested. The addition of octylamine to the reaction medium increases the velocity of DMA N-oxidation, but it apparently doesnot affect the binding of DMA to the oxidase. The concentration of DMA required to half-saturate the oxidase, calculated from the data in Fig. 4, is 2.9 X W5 M both in the presence and absenceof n-octylamine. Activation of the DMA N-oxidase by n-otcylamine is also observed in microsomes
MICROSOMAL
ACTIVATION
OF THE
OF MICROSOMES
TABLE DMA
n-Octylamine Cadaverine Phenethylamine Amphetamine Mescaline Aniline p-Chloroaniline
IV
123
III
TABLE
N-OXIDASE
-*ND THE PURIFIED BY PRIMARY AMINES
Addition to the reaction medium Compound
OXIDASE
ACTIVITY’
EFFECT
OF VARIOUS DMA OXID~SE
ENZYME
PURIFIED
@a - vc)I(vJ x 1oob
Concentration (mrd)
Microsomes
Isolated amine oxidase
1.0 2.0 2.0 2.0 1.0 5.0 0.5c 1.0 5.0 1.0
87 115 0 22 30 82 22 0 0 0
85 98 0 16 26 74 17 0 0 0
6 DMA N-oxidase activities were calculated from the rate of DMA N-oxide formation at pH 8.4. The composition of the basic reaction media was as described under Methods. The concentration of DMA was 2.5 mu. b va = DMA N-oxidase activity in the presence of the primary amine. v0 = DMA N-oxidase activity in the absence of primary amine. c This is the highest concentration soluble in the reaction medium.
Compound Carbon monoxide” Nitrogenb SKF-525A Cyanide Azide 1-(1-Naphthyl)-2thiourea Catalase Erythrocuprein
IV COMPOUNDS
ACTIVITY
ON THE OF THE
ENZYMES
Concentration Icol/[ozl = 4 in gas phase 100% in gas phase 0.5 rnM 1.0 rnM 1.0 rnM 0.05 rnM 0.50 rnM 1 mg/ml 0.1 mg/ml
$?~~mn 0.0 99 0.0 0.0 0.0
33 92 0.0 0.0
mThe assays were carried out as described in Methods. The concentration of DMA was 2.5 rnM; the reaction time was 2 min. b The reaction vessels were gassed during the 5-min temperature equilibration period. Enzyme, followed by DMA, was added to start the reaction; and the vessels were sealed during the reaction. DMA added prior to temperature equilibration is removed almost quantitatively by a gas flow sufficient to equilibrate the reaction medium.
tested that either have no effect or that inhibit the DMA N-oxidase at high concentrations are octanol, cyclohexanol, nonyl as shown by the information in Table III. As observed with the purified oxidase, n- sulfate, and hexanoic and octanoic acids. Inhibitors. As indicated by the data in octylamine increases the rate of DMA N-oxidation catalyzed by the hepatic Table IV, the amine oxidase is insensitive to microsomes. The increase in pig liver carbon monoxide and SKF-525A, two commicrosomal DMA N-oxidase produced by pounds frequently used to inhibit micro2 mM n-octylamine is usually 2-fold, but somal mixed-function oxidases. The amine 1.3- to 3.0-fold activations have been ob- oxidase activity of the enzyme is also comtained with different preparations of pig pletely insensitive to cyanide and azide even liver microsomes. This is in contrast to the at concentrations 10 times greater than purified oxidase where 2 mu octylamine those listed in the table. The residual cataconsistently stimulates the N-oxidation of lase activity always present in the oxidase is, DMA 1.9-fold. however, inhibited almost 100% by 1 mu Other primary alkylamines will also azide or cyanide. Both of these compounds stimulate the DMA N-oxidase activity of inhibit the oxidase-catalyzed peroxidation of both microsomes and the purified oxidase. methanol and ethanol by hydrogen peroxide. Phenethylamine, amphetamine, and mesca- This activity is probably due to contaminaline are not as effective as n-octylamine; but tion of the purified oxidase by catalase and stimulation of the DMA N-oxidase by these does not appear to be a property of the amines is always observed. Neither ca- amine oxidase. Neither catalase nor erythrodaverine (a diamine) nor aniline affects the cuprein, added in large excessto the reaction DMA N-oxidase activity of microsomes or medium, affects the amine oxidase activity of the purified enzyme. Other compounds of the purified enzyme.
124
ZIEGLER
AND
Of all the compounds tested, l-(lnaphthyl)-2-thiourea appears to be the most selective inhibitor of the amine oxidase, even though this compound would be more accurately described as a substrate with a high affinity for the oxidase but with a very low turnover rate. The more detailed studies of Fok (23) in this laboratory suggest that I-(1-naphthyl)-2-thiourea is competitive with the amine substrate with an apparent Ki of 3 X 10m6M. This compound alone will also slightly stimulate the endogenous NADPH oxidase activity of the purified oxidase, and the naphthylthiourea appears to be consumed during the reaction. This latter interpretation is based on the observation that 0.010-0.015 mM naphthylthiourea, preincubated with NADPH and the oxidase, loses its ability to inhibit the oxidation of DMA. If either NADPH or enzyme is omitted during preincubation, the compound retains its ability to inhibit the oxidation of DMA by the oxidase. It would is appear that 1-(1-naphthyl)-2-thiourea enzymically oxidized to products that do not inhibit the oxidase; but oxidation products of this compound have not, as yet, been identified. While the data are not included in Table IV, 1-(1-naphthyl)-2-thiourea also inhibits the N-oxidation of amines catalyzed by pig or rat hepatic microsomes; but this compound has very little effect on the microsomal-catalyzed oxidative N-demethylation of aminopyrine or propoxyphene. Concentrations of 1-(1-naphthyl)-2-thiourea that inhibit the microsomal N-oxidase activity 80% inhibit the demethylation of the latter two substrates less than 10%. While more work on the effects of 1-(1-naphthyl)-2thiourea on microsomal oxidases will be necessary, these studies indicate that this compound may be a useful t,ool for distinguishing between oxidations catalyzed by the amine oxidase or by other microsomal oxidases. DISCUSSION
The purified oxidase has not been fully characterized as to its amine substrate specificity or its physical properties or ultrastructure. An earlier report (14) with a par-
MITCHELL
tially purified preparation demonstrated that, this oxidase catalyzed the N-oxidation of a large number of synthetic drugs and naturally occuring alkaloids. The highly purified oxidase described in this report N-oxidizes all of the compounds listed in the earlier rcport at increased rates proportional to the increase in the purity of the oxidase. The studies of Sakurai, currently in progress in this laboratory, indicate that the isolated oxidase has a molecular weight close t.o 500,000 and is composed of similar, if not identical, flavoprotein subunits. The precise function and organization of the flavoproteins in the active complex remain to be determined. Earlier studies both in vitro and in viva indicated that cyt. P-450 was not a component of the hepatic mixed-function oxidase that catalyzes the N-oxidation of set- and tertamines. Both carbon monoxide and SKF525A, widely used inhibitors of the microsomal-cyt. P-450 oxidase system, do not inhibit the N-oxidation of amines in vitro (6) ; and in rats the latter compound produced an increased urinary excretion of amine oxides when injected simultaneously with simple dimethylarylor dimethylalkylamines (24). Uehleke et al. (25) have reported that hepatic microsomes, irradiated with uv light to inactivate cyt. P-450, still catalyzed the N-oxidation of secondary and tertiary amines, again indicating that cyt. P-450 was not required for the N-oxidation of these amines. These conclusions are supported by the work presented in this paper. The isolation of a mixed-function amine oxidase free from cytochromes demonstrates that, a microsomal electron transport system is not required for the N-oxidation of set- or ted-amines. A recent direct comparison (26) of the mixed-function amine oxidase and NADPH-cyt. c reductase, both isolated from pig liver microsomes, demonstartes unequivocally that the two flavoproteins are separate enzymes with distinct physical and catalyt,ic properties. Since in microsomes NADPH-cyt. c reductase catalyzes the reduction of cyt. P-450, it would appear that hepatic microsomes contain separate enzyme systems catalyzing the N-oxidat,ion and oxidative-N-dealkylation of many N, Ndimethylalkylor N, N-dimethylaryl-amines.
MICROSOMAL
However, not all microsomal-catalyzed N-oxidations can be attributed to the mixedfunction amine oxidase. The purified amine oxidase does not catalyze the N-oxidation of aniline or of p-chloroaniline amines that are known to be N-oxidized by hepatic microsomes (12). The recent report by Uekleke et al. (25) also suggeststhat different microsomal enzymes were responsible for the cat,:Jytic N-oxidations of primary amines an*f tertiary amines. With the few exceptions described in a recent report from this laboratory (27), it would appear that the N-oxidations of primary amines are catalyzed by a microsomal oxidase different from the mixed-function amine oxidase described in this report. Primary alkylamines, however, do interact with the amine oxidase as shown by the data in Figs. 3 and 4, and Table III. While these amines are not oxidized, they increase the DMA oxidase activity of the enzyme; and the activation of this activity by primary amines is also consistently observed with the microsomal-bound enzyme. Activation of this microsomal oxidase by alkylamines appears to be an inherent property of the enzyme and may reflect a mechanism that controls the activity of this oxidase in vivo. ACKNOWLEDGMENTS We are indebted to Dr. Johnston at Sout,hwestern Medical School, University of Texas at Dallas, for the analysis of the lipids associated with the purified amine oxidase. We also are indebted to Dr. Yukihiko Sakurai for his assistance in perfecting the zone electrophoresis step in the isolation of the oxidase. REFERENCES E. W., AND 1. MACHINIST, J. M., DEHNER, ZIEGLER, D. M. (1968) Arch. &o&m. Biophys. 126, 858. 2. BAKER, J. R., STRUEMPLER, A., AND CHAYKIN, S. (1963) Biochim. Biophys. Acta 71,58. 3. BAKER, J., AND CHAYKIN, S. (1962) J. Biol. Chem. 937, 1309.
OXIDASE
IV
125
4. MASON, H. S. (1957) Science 126, 1185. 5. CONNEY, A. H. (1967) Pharmacol. Rev. 19,317. 6. ZIEGLER, D. M., AND PETTIT, F. H. (1966) Biochemistry 6, 2932. 7. HOPPE-SEYLER, F. A. (1934) Ber. Gesamte Physiol. Exp. Pharmakol. 81, 392. 8. LINTZEL, W. (1934) Biochem. Z. 273,243. R., PHILLIPS, A., TSAI, I., 9. KUNTZMAN, KLUTCH, A., AND BURNS, J. J. (1967) J. Pharmacol. Exp. Ther. 166, 337. 10. BICKEL, M. H. (1969) Pharmacol. Rev. 21, 325. 11. BECKETT, A. H., AND HEWICK, D. S. (1967) J. Pharm. Pharmacol. 19, 134. 12. KIESE, M. (1966) Pharmacol. Rev. 18, 1091. 13. HLAVICA, P., AND KIESE, M. (1969) B&hem. Pharmacol. 18, 1501. 14. ZIEGLER, D. M., MITCHELL, C. H., AND JOLLOW, D. (1969) in Microsomes and Drug
15. 16. 17. 18.
Oxidations (J. R. Gillette, A. H. Conney, G. J. Cosmides, R. W. Estabrook, J. R. Fouts, and G. J. Mannering, eds.), p. 173. Academic Press, New York. DAS, M. L., AND ZIEGLER, D. M. (1970) Arch. Biochem. Biophys. 140, 300. ZIEGLER, D. M., AND PETTIT, F. H. (1964) Biochem. Biophys. Res. Commun. 16, 188. FOK, A. K., AND ZIEGLER, D. M. (1970) Biothem. Biophys. Res. Commun. 41, 534. BROWN, J., AND JOHNSTON, J. M. (1962) J. Lipid
Res. 3, 480. J., AND PETERSON, N. F. (1965) J. Lipid Res. 7, 455. HAUSCHILD-ROGAT, P., AND SMITH, I. (1968) in Chromatographic and Electrophoretic
19. PARKER, 20.
21. 22. 23. 24. 25.
26.
27.
Techniques. (Ivor Smith, ed.), 2nd edition, p. 475. Wiley (Interscience), New York. DAVIS, B. J. (1964) Ann. N. Y. Acad. Sci. 121, 321. MITCHELL, C. H., AND ZIEGLER, D. M. (1969) Anal. Biochem. 28, 261. FOK, A. K. (1971) Ph.D. Thesis, Department of Chemistry, University of Texas. DEHNER, E. W., MACHINIST, J. M., AND ZIEGLER, D. M. (1968) Life Sci. 7, 1135. UEHLEKE, H., SCHN~TGER, F., AND HELLMER, K. H. (1970) Hoppe Seyler Z. Physiol. Chem. 361, 1475. MASTERS, B. S. S., AND ZIEGLER, D. M. (1971) Arch. Biochem. Biophys. 146, 368. ZIEGLER, D. M., POULSEN, L. L., AND MCKEE, E. M. (1971) Xenobiotica 1,4.
OF
BIOCHEMISTRY
Microsomal
AND
Oxidase Oxidase DANIEL
Clayton
Foundation
116-125
BIOPHYSICS
160,
IV:
Properties
Isolated M. ZIEGLER2
(1972)
of a Mixed-Function
from
Pig
Liver
CAROLYN
AND
Microsomes’ H. MITCHELL3
Biochemical Institute and the Department of of Texas at Austin, Austin, Texas Received
December
22, 1971;
accepted
Amine
February
Chemistry,
The
University
3, 1972
A flavoprotein-oxidase isolated from pig hepatic microsomes catalyzes the NADPHand oxygen-dependent N-oxidation of a variety of lipid-soluble set- and tert-amines. The set-amines are oxidized to the hydroxylamines; and the tert-amines, to the amine oxides. With one or two exceptions the isolated oxidase does not catalyze the oxidation of primary amines. However, primary alkylamines added to the reaction vessel increase the set- and tert-amine N-oxidase activity of both the isolated enzyme and microsomes. The isolated oxidase contains 14 nmoles FAD and about 0.02 mg lipid/mg protein. The oxidase appears to be free from cytochromes but is contaminated with very small, but detectable, amounts of a hemoprotein that corresponds to catalase, both spectrally and in enzymic properties. The isolated enzyme is free from NADPH-cyt. c reductase, diaphorase, and monoamine oxidase activities.
An earlier report (1) demonstrated that hepatic tissues from vertebrates contained an oxidase that catalyzed the oxidation of of DMA4 to the amine oxide. While this activity could be detected in all vertebrates tested, the DMA N-oxidase activity was consistently highest in hepatic tissue from pigs. The speciesand organ distribution of DMA N-oxidase activity resembled the distribution of the trimethylamine N-oxidase described by Baker et al. (2). Both DMA N-oxidase and the trimethylamine Noxidase require NADPH for activity, and both activities are concentrated in the microsomal fraction of liver homogenates. ‘This work was supported in part by U. S. Public Health Service Grant GM 12360. ZWork carried out during tenure of a U. S. Public Health Service Career Development Award (l-K3GM-25,990). 3California Institute of Technology, Pasadena, CA. ‘Abbreviations used are: DMA, N,N-dimethylaniline ; DMOA, N, N-dimethyl-n-octylamine; SKF-525A, Diethylaminoethanol ester of diphenylpropylacetate. 116 Copyright
@ 1972 by Academic
Press,
Inc.
Baker and Chaykin (3) have also shown that the oxygen atom incorporated into trimethylamine is derived from molecular oxygen. The enzyme catalyzing the Noxidation of DMA appeared to be of the type classified as mixed-function oxidases (4). Unlike other widely studied hepatic microsomal mixed-function oxidases (5)) the DMA N-oxidase was reported to be insensitive to both carbon monoxide and SKF-525A(6). This and other evidence suggested that cyt. P-450 was not an obligatory component of the microsomal DMA Noxidase system. Both in vitro and in vivo N-oxidation of other tertiary amines has been demonstrated in many vertebrates including man (1, 2,7-g). While N-oxides are generally considered minor metabolites of amine metabolism, the rapid N-oxidation of certain tertiary amine drugs has been reperted (cf. review by Bickel, 10); and Beckett and Hewick (11) have shown in vitro that amine oxides are the major oxidation products of the phenothiazine drugs.
MICROSOMAL
The N-hydroxylation of secondary amines has also been observed in vitro, and the extensive studies of Kiese and his associates (12, 13) have shown that the organ and subcellular distribution and the cofactor requirements of the secondary amine Nhydroxylase are virtually identical with those of the tertiary amine N-oxidase. Their studies support the interpretation that the same enzyme catalyzes the N-oxidation of both secondary and tertiary amines. This report describes the isolation from pig liver microsomes of a flavoproteinoxidase that catalyzes the NADPH- and oxygen-dependent N-oxidation of secondary and tertiary amines. Some of the properties of partially purified preparations of this oxidase were described in an earlier report (14). EXPERIMENTAL
aminesubstrates,obtainedfrom commercialsources,were purified by gas-liquid Materials.
The
chromatography and then stored at -15°C. As needed, aliquots of the pure amines were dissolved in equivalent amounts of 0.1 M HCl. Solutions of the aniline derivatives were prepared daily. Aqueous solutions of the other amines are more stable and were stored at - 15°C for several weeks without undergoing any apparent change. Ultrapure sucrose and ammonium sulfate were obtained from Mann Research Lab; Alphacel and protamine sulfate, from Nutritional Biochem. Corp. ; NADP+, NADPH, Sephadex G-100-120, DEAE-cellulose, Triton-X-45, Triton X-100, and Triton-X-102, from Sigma Chemical Company. In addition, samples of all the detergents used in this study were initally supplied by R,ohm, Haas and Company, Philadelphia. Ten-per cent solutions of each of the Triton detergents were prepared by dissolving 100 g of the detergent in water to a final volume of 1 liter. The solutions were stored at 0-5°C. Even though a mixture of Triton-X-102 and Triton-X-45 was used to extract the oxidase, these two detergents had to be stored as separate solutions. Equal volumes of the two detergent solutions were mixed just prior to use. The saturated “acid’‘-ammonium sulfate solution was prepared by dropwise addition of concentrated sulfuric acid to a solution saturated at room temperature with ammonium sulfate. Sufficient sulfuric acid was added to bring the pH of the solution to 4.8 f 0.1 at 25’C after it was diluted lo-fold with water. The acid-ammonium sulfate solution was stored at O-5’% for at least 1 day before use.
OXIDASE
IV
117
The pretreatment and conditions for storing DEAE-cellulose, Sephadex, and cellulose were as follows: The DEAE-cellulose was suspended in about 10 vol of 0.5 M HCl and then washedwith water until the effluent was free from chloride ions. The washed ion-exchange cellulose was resuspended in about 10 vol 0.03 M glycine and stored at 0-5°C. Fifteen grams of Sephadex G-100-120 suspended in 2 liters of water were heated in a steam cabinet for 8 hr. Before cooling, HCl was added to the hydrated Sephadex to a final concentration of 0.1 M; and the suspension was heated to boiling for 5 min. The mixture was rapidly cooled on ice, and the Sephadex was washed with cold water until free from chloride ions. The gel, suspended in 5 vol 0.05 M glycine, can be stored for several months at 0-5°C. The cellulose (Alphacel) was treated three to four times (alternately) with 0.02 M HCl and 0.02 M KOH to remove extraneous material. The cellulose was collected by filtration, washed with a large excess of water to remove the last traces of KOH (pH of the filtrate about 6), resuspended in water, and stored in the refrigerator until needed. Pig liver microsomes were isolated by the method described earlier (6). The liver tissue was obtained almost exclusively from old sows since, as shown with rats (15), the hepatic N-oxidase activity is higher per milligram of protein in tissue from older animals. Methods. Unless otherwise indicated, enzyme assays were carried out in open lo-ml Erlenmeyer flasks as described earlier (16). The reaction medium routinely contained 0.5 mm NADP+; 5.0 rnM magnesium chloride; 5 mM isocitrate; isocitrate dehydrogenase (sufficient to reduce 1 pmole NADP+/min/ml); and 0.1 M alanine-0.02 M pyrophosphate buffer, pH 8.4, at 38°C. After a 5-min temperature equilibration the reaction was started by the addition of enzyme. The amine oxidation products were measured as described earlier (1517). Protein and acid-extractable flavin were measured by standard methods (cf. Ref. 15). Lipids extracted by the Folch procedure from the purified oxidase were assayed for neutral lipids by the method of Brown and Johnston (18). The phospholipids were separated by tic, and the phosphate concentration of the individual phospholipids was measured by the method of Parker and Peterson (19). Isolation of the mixed-function amine N-oxidase. All steps in the procedure were carried out at O-5’ C. Microsomes stored in 0.25 M sucrose at - 15’C for no longer than 1 month were thawed, diluted to approximately 20 mg protein/ml, and adjusted to pH 5.8 by dropwise addition of 1 M acetic acid. The suspension was immediately centrifuged at 30,000 rpm for 10 min. The residue was collected and re-
118
ZIEGLER
PURIFIC.~TION
OF THE MIXED-FUNCTION
AND
MITCHELL
TABLE AMINE
I OXIDASE
Volume (ml)
Fraction
FROM Protein (w)
-_ ..Microsomes 3550%
Ammonium
sulfate
Acid-ammonium
sulfate
After
DEANE-cellulose
After After
Sephadex G-100-129 electrophoresis
(1 The assays was 2.5 mM; the b The values tions that have
chromatography chromatography
400
9600
120
830
16
150
8
118
2 3
75 26
PIG
LIVER
nmoles DMA oxidized min/mg
protein
23.0 (16-38)” 160 (99-210) . 520 (36&600) (47E20)
Percent of original activity 100 60 35 33
890 1,049
were carried out at 38°C as described in Methods. The initial reaction time was 3 min. listed in parentheses are the lowest and highest specific activities been successfully carried through the succeeding steps.
suspended in water to approximately 20 mg protein/ml and then was adjusted to pH 8.0 with dilute KOH. Four milliliters of 2.5 M guanidine hydrochloride were added for each 100 ml of the suspension, and the mixture wasthoroughly homogenized in a glass homogenizer fitted with a Teflon pestle. The aggregated particles were sedimented by centrifugation at 30,000 rpm for 15 min. The residue was resuspended in water and homogenized. The volume was adjusted to give a protein concentration of 29 f 1.0 mg/ml. The oxidase was extracted from the particles by adding, for each 100 ml of the suspension, 9 ml of the Triton X-45Triton X-102 solution (see Materials section). After stirring for 1 hr, 100 mg protamine sulfate were added for each 100 ml of the suspension. The correct amount of protamine sulfate was routinely dissolved in a minimum volume of water at 38°C and slowly added to the microsomal suspension with rapid stirring. The detergent-insoluble material was removed by centrifugation at 30,900 rpm in a Spinco Model L centrifuge for 1 hr. The clear, yellow-brown supernatant fraction was collected and immediately fractionated with ammonium sulfate. The fraction that precipitated upon the addition of 19.4 g ammonium sulfate for each 100 ml of solution was removed by centrifugation. This fraction collected as an oily pehicle at the top of t.he centrifuge tube and was removed and discarded. The soluble fraction was, if necessary, carefully filtered through glass wool to remove any remaining insoluble material; and an additional 8.7 g ammonium sulfate were added for each 100 ml of solution to precipitate the oxidase. The precipitate was collected by centrifugation and re-
MICROSOMEP
30 12
concentration of different
of DMA prepara-
dissolved in water to a final protein concentration of not more than 8 mg/ml. This fraction is referred to in Table I as the 35-5070 ammonium sulfate fraction. One-tenth volume of 1 M sodium acetate, pH 4.8, was added to the 35-5Oyc ammonium sulfate fraction; and the preparation was refractionated by adding the saturated acid-ammonium sulfate solution (see Materials section). Fractions precipitating at O-35,35-40,40-45, 4550% saturation were collected by centrifugation and redissolved in 0.05 M glycine, pH 6.2-6.6. The 0-35yc saturation fraction usually contained little or no amine oxidase activity and was discarded. The enzyme was usually most concentrated in the 4045% ammonium sulfate fraction while the other two fractions usually contained significant amounts of the oxidase. The latter two fractions (3540$& and 4550%) were combined, and sufficient saturated acid-ammonium sulfate was added to produce a faint turbidity. After stirring for 30 min, the precipitate was removed by centrifugation and discarded. The oxidase in the supernatant fraction was precipitated with ammonium sulfate (about 450/o saturation), collected by centrifugation, and combined with the initial 40-45’% saturation fraction. The combined fractions are referred to as the acid-ammonium sulfate fraction in Table I. Triton-X-100 (0.1 mg/ml protein) was added to the acid-ammonium sulfate fraction before it was carried through the final steps. After adding the detergent, the preparation was dialyzed with internal and external stirring against 20 vol water for 90 min. The water was changed every halfhour. The dialyzed preparation, in 10 ml or less,
MICROSOMAL was passed through a column (20 X 2.5 cm) of DEAE-cellulose that had been equilibrated with several column volumes of a solution containing 0.03 M glycine and 0.03 M histidine, pH 6.3. The oxidase, an intense yellow band, was washed through the column with 0.03 M histidine buffer, pH 6.3. The eluate containing the oxidase was reduced in volume to about 2 ml by pressure dialysis in an Amicon Diaflo Cell (Model 52) fitted with an XM-100 membrane. The concentrated fraction was immediately placed on a column (20 X 2.5 cm) of Sephadex G-100-120 equilibrated with 0.03 M glytine. Four-milliliter fractions were collected at a flow rate not greater than 0.5 ml/min. The fractions containing the oxidase were combined and reduced in volume to 1.5-2 ml by pressure dialysis. The final step in the isolation of the oxidase was carried out by zone electrophoresis in a unit similar to that described by Hauschild-Rogat and Smith (20) for preparative disc electrophoresis, except that the polyacrylamide gel was replaced with cellulose. The buffers used were the same as those specified by Davis (21) for polyacrylamide gel-disc electrophoresis. A maximum of 2 ml of each Sephadex G-100-120 fraction, containing Tris buffer, pH 6.7, was carried through the electrophoresis step at one time. In 5 hr at 350 V and 10 mA, the oxidase migrates 4-6 cm as a narrow, intensely yellow band. When the enzyme was about 2-3 mm from the bottom of the cellulose column, the electrodes were removed; and the oxidase was eluted with the electrode buffer at a flow rate not greater than 0.1 ml/min. The yellow fraction, in a volume of 2-3 ml, was collected and concentrated almost to dryness by pressure dialysis. The enzyme was taken up in 3-5 ml of 0.05 M histidine, pH 6.6-7.0, and stored at 0-3°C. Stability of the amine oxidase during isolation. The oxidase is quite unstable immediately after it is extracted from the particles, and all of the steps through the 35-500/, ammonium sulfate fraction must be carried out as rapidly as possible. The 35-50y0 ammonium sulfate fraction, if diluted to a protein concentration between 5-8 mg/ml, can be stored overnight at 0-5°C with very little loss of activity. If this fraction is stored at a higher protein concentration or for longer periods of time, progressive inactivation of the enzyme is frequently observed. In contrast, the oxidase in the acid-ammonium sulfate fraction is quite stable and can be stored at -15°C for several months with little or no loss of activity.After Sephadex chromatography the oxidase remains fully active when stored at 0-3°C for up to 2 weeks, but 20-40~0 of the activity is lost every time the preparation is frozen and thawed. Inactivation of the purified oxidase by freezing and thawing can be retarded, but not completely prevented, by the addition of 0.1 mg
OXIDASE
119
IV
Triton-X-lOO/mg enzyme. The most stable form of the oxidase is found in the acid-ammonium sulfate fraction, and the preparations are routinely stored at this stage. As needed, aliquots of this fraction are carried through the final steps in the isolation. RESULTS
The concentration of the oxidase varies widely in different preparations of pig liver microsomes (6). As a result the yield and concentration of the oxidase obtained during isolation were also extremely variable. If the initial specific activity of the microsomes is less than 15 nmoles DMA N-oxidized/ min/mg protein, the yield, st.ability, and purity of the oxidase are very erratic. Preparations of the oxidase at the highest level of purification are consistently obtained from microsomal fractions with an initial DMA N-oxidase specific activity of 18 or greater, and the concentration of the oxidase/mg protein obtained after the different steps is as indicated in Table I. The concentration of the enzyme after passage through DEAE-cellulose is not great and is somewhat variable, but this step is necessary to remove the last traces of cyt. bg. Any cyt. bgnot removed at this point will concentrate with the oxidase during chromatography on Sephadex. Both the pH and ionic strength of the amino acid buffer during DEAE-cellulose chromatography are critical. At a more neutral pH or lower ionic strength, the oxidase is irreversibly adsorbed on the ion-exchange cellulose. Irreversible binding of the enzyme on Sephadex G-100-120 does not occur; and at the low ionic strength used, virtually all of the purification of the oxidase may be attributed to the ion-exchange properties (enhanced by heating in dilute acid; cf. Methods section) of the Sephadex. In the presenceof 0.1 M KCl, very little purification of the oxidase is obtained by chromatography on Sephadex G-100-120. Composition of the oxidase. The purified oxidase contains 14 + 0.2 nmoles of flavin/ mg protein. All of the flavin can be ext’racted with acid and appears to be identical with FAD both spectrally and in its ability to activate the apo-D-amino acid oxidase. The amine oxidase contains about 2% lipid by weight. Analysis of the lipids extracted by
120
ZIEGLER
AND MITCHELL
the E’olch procedure from 23 mg of the purified oxidase showed the presence of the following in micrograms per milligram protein: phosphatidyl choline, 14.7; lysophosphatidyl choline, 2.7; free fatty acids, 3.9; unidentified phospholipids, 0.9; and small but detectable amounts of cholesterol. The purified oxidase appears to be entirely free from the detergents used to extract the enzyme from the particles. The characteristic uv absorption bands of the Tritons cannot be detected in dried acetone-extract’s of the oxidase redissolved in methanol or wat,er. Tritons added to the purified enzyme can be readily extracted and identified by this procedure. The oxidase does not contain detectable amounts of iron, copper, or cytochromes. The purified oxidase is also free from detectable NADH or NADPH diaphorase, cyt. c reductase, and monoamine oxidase activities. The purified preparation contains small but detectable amounts of catalase activity. Spectra. The spectrum of the oxidized flavoprotein (Fig. 1) is similar to that of other flavoproteins. The 260 nm/460 nm absorption ratio is greater than that of most flavoproteins. The absorption of the oxidized enzyme at 410 nm is also greater than expected for a pure flavoprotein. The increased absorption in this region of the spectrum may be due to the presence of small amounts of hemoprotein contaminant, probably catalase. Anaerobically, both the 380 and 450 absorption bands of the oxidase are completely reduced by NADPH; but in the presence of oxygen, NADPH produces no perceptible change in the spectrum of the oxidized enzyme. NSDPH oxidase activity. The isolated oxidase catalyzes the oxidation by oxygen of 90-110 nmoles of NADPH/min/mg protein at 38°C. A number of observations suggest that this activity is an inherent property of the oxidase and is not due to a contaminating NADPH-dependent enzyme. The pH optimum (8.4) for the NADPHoxidase activity is identical with the pH opt’imum for the N-oxidase activity of the enzyme. The ratio of the DMA-N-oxidase/ NADPH-oxidase activities remains relatively constant (10/l) during the last four
320 WAVELENGTH
400
460
560
my
FIG. 1. Direct spectraof the purified amineoxidase. The concentration of the enzyme is 0.8 mg protein/ml in 0.1 M glycine-O.02 M pyrophosphate, pH 8.4. -, oxidized enzyme; * * * * *, enzyme anaerobically reduced by 0.1 mM NADPH; and -----, enzyme reduced with NazS204. The spectrum of the NADPH-reduced enzyme was recorded after deaeration of the sample with nitrogen followed by addition of the isocitrate dehydrogenase-NADPH generating system. The same concentrations of NADPH and the generating system were added to the reference cuvette to minimize interference by the 340-nm band of NADPH with the spectrum of the reduced enzyme.
steps in the isolation of the oxidase. Attempts to differentiate between the two activities by pretreating the oxidase with heat detergents, mecurials, and proteolytic enzymes have been unsuccessful. Both activities are destroyed at the same rate. In the absence of an amine substrate the molar ratio of NADPH oxidized/Oz consumed, measured polarographically, is 2.0 (Poulsen, unpublished experiments this laboratory). Since the preparation also contains small quantities of catalase, these data suggest that hydrogen peroxide is produced during the reoxidation of the flavoprotein by oxygen. In the presence of saturating levels of added catalase, the purified oxidase will catalyze the peroxidation of methanol at about the same rate as that for NADPH oxidation, indicating that hydrogen peroxide is produced during the oxidation of NADPH.
MICROSOMAL
N-OXIDATION
TABLE OF AMINES PURIFIED AMINE
Substrate
II C.~T.~LYZED 0x1~~~~
nmoles oxidation product/min/mg protein N-Oxide
Tertiary amines Trimethylamine N, N-dimethylbutylamine N,N-dimethylhexylamine N, N-dimethylcyclohexylamine N,N-dimethyloctylamine N, N -dimethylnonylamine N,N-dimethyldodexylamineb N, N -dimethylaniline N, N -diethylaniline N-methylmorpholine N-ethylmorpholine Secondary amines N-methylaniline N-ethylaniline N-methyloctylamine
BY THE
Hydroxylamine
260 546 552 516
-
1650 1760 360
-
910 712 378 284
-
-
507 380 1020
a The fraction obtained after Sephadex G-lOO120 chromatography. Except for N-ethylmorpholine, the rates listed for all amine substrates were calculated from the rate of oxidation-product formation at 38°C in 2 min with 2.5 mnn amine substrate and 0.1-0.3 mg enzyme/ml. The rate for N-ethylmorpholine is based on the substratedependent oxidation of NADPH. b Concentration of this substrate was 0.5 mu.
Amine oxidation products. In addition to DMA, the purified amine oxidase catalyzes the N-oxidation of a number of other secondary and tertiary amines (Table II). The tertiary amines are oxidized to the corresponding amine oxides; and the secondary amines, to hydroxylamines. As shown earlier (22) with hepatic microsomes, the N, N-dimethyl-n-alkylamines with S-12 carbons in the side chain are N-oxidized at the fastest rates. The size, but not the arrangement, of the alkyl side cha,in appears to determine the rate at which the oxidase catalyzes N-oxidation of the substrate. For example, the N, N-dimethyl derivatives of n-hexyland cyclohexyl-amines are Noxidized at about the same rate. Amines
OXIDASE
IV
121
with N-ethyl groups are oxidized at slower rates than the corresponding N-methyl compounds (cf. derivatives of morpholine and aniline in Table 11). The secondary N-methyl amines are oxidized at about N the rate of the corresponding N, N-dimethyl tertiary amines. While only a limited number of secondary amines have been extensively studied, the effect of the side chain on the oxidation rate appears to be similar to that observed with the tertiary amines. Hydroxylamines appear to be the only oxidation products obtained from secondary amines. An earlier report indicated that a partially purified preparation of the amine oxidase catalyzed the oxidative N-demethylation of secondary amines (14). This interpretation was based on t’he observations that substantial amounts of formaldehyde can be detected in deproteinized aliquots of the reaction media containing the secondary amine substrates and the oxidase and t.hat the formation of formaldehyde was dependent upon substrate concentration and incubation time. However, it would appear that the formaldehyde is an aciddegradation product of the enzymically formed Nmethylalkylhydroxylamines. If aliquots of the reaction mixture are extracted with ether to remove hydroxylamines before deproteinizing with trichloroacetic acid, formaldehyde cannot, be detected in the aqueous phase. Formaldehyde added to the reaction vessels, with or without hydroxylamine, can be qualitatively recovered in the aqueous phase after repeated extraction with ether. The purified oxidase does not catalyze the oxidation of most primary amines or of secondary or tertiary amines with polar groups near the nitrogen. Oxidation of the N-methyl derivatives of acetanilide, formanilide, and nicotinamide could not. be detected. Stoichiometry. As indicated by the experiments summarized in Fig. 2, 1 mole NADPH is required for each mole of substrate Noxidized. The ratio of the rate of DMOAdependent NADPH oxidation to the rate of DMOA N-oxide formation is 1: 1. However, the rate of DMA N-oxide formation is somewhat greater than the rate of DMAdependent NADPH oxidation and is equal
122
ZIEGLER
AND MITCHELL
I 0 0
I
2 TlME
3
4
I 2
I 4
t 6
I
5 I/DMOAlmM)-'
(mill)
FIG. 2. Rates of NADPH oxidation and amine oxide formation. The reactions were carried out at 38°C in 3.0~ml cuvettes in a Zeiss PM-&II spectrophotometer. NADPH concentrations were measured at 340 nm, and aliquots of the reaction mixtures were withdrawn and analyzed for amine oxides at the times indicated. A-A, rate of DMOA-N-oxide formation. O-O, rate of DMOAdependent NADPH oxidation. A-A, rate of DMA-N-oxide formation. a-0, rate of DMAdependent NADPH oxidation. The initial concentrations of NADPH, DMOA, and DMA were 0.2, 3.0, and 1.5 mM, respectively.
to, or slightly less than, the total rate of NADPH oxidation. Thus, DMA, in contrast to DMOA, appears to block the endogenous NADPH oxidase activity of the isolated oxidase. E$ect of primary amines on activity. Double reciprocal plots of DMOA concentration versus reaction rate are shown in Fig. 3. The points do not fall on a straight line, and a pronounced concave curve is obtained over the range of DMOA concentrations tested. High concentrations of this substrate apparently activate the oxidase, and this activation is quite evident at DMOA concentrations above 1 mu. However, amines that are not substrates for the oxidase can also activate the enzyme. As shown in Fig. 3, the enzymic rate at low DMOA concentrations is almost doubled in the presence of 5 mu n-octylamine. At the highest concentration of DMOA tested, the addition of octylamine did not increase the oxidation rate of DMOA; and both curves in Fig. 3 appear to intersect the y axis at about the same point. In cont’rast to the alkylamines, the aniline
FIG. 3. l/DMOA vs. l/v, where v is the rate of DMOA-N-oxide formation/min/mg protein at pH 8.4 and 38°C. O-0, DMOA. wm, DMOA in the presence of 5.0 lllM n-octylamine.
‘,“I 1.0
-
z 0.5-
-40
-30
-20
-10
0
10
20
-I
l/DMA
(mM1
FIQ. 4. l/DMA vs. l/v, where v is the rate of DMA-N-oxide formation/min/mg protein at pH 8.4 and 38°C. +-@, DMA. H, DMA in the presence of 1.0 rnM n-octylamine.
derivatives do not activate the oxidase. As shown in Fig. 4, double reciprocal plots of DMA concentration versus rate yield straight lines. The addition of 1.0 mu n-octylamine doubles the rate of DMA N-oxidation over the whole range of DMA concentrations tested. The addition of octylamine to the reaction medium increases the velocity of DMA N-oxidation, but it apparently doesnot affect the binding of DMA to the oxidase. The concentration of DMA required to half-saturate the oxidase, calculated from the data in Fig. 4, is 2.9 X W5 M both in the presence and absenceof n-octylamine. Activation of the DMA N-oxidase by n-otcylamine is also observed in microsomes
MICROSOMAL
ACTIVATION
OF THE
OF MICROSOMES
TABLE DMA
n-Octylamine Cadaverine Phenethylamine Amphetamine Mescaline Aniline p-Chloroaniline
IV
123
III
TABLE
N-OXIDASE
-*ND THE PURIFIED BY PRIMARY AMINES
Addition to the reaction medium Compound
OXIDASE
ACTIVITY’
EFFECT
OF VARIOUS DMA OXID~SE
ENZYME
PURIFIED
@a - vc)I(vJ x 1oob
Concentration (mrd)
Microsomes
Isolated amine oxidase
1.0 2.0 2.0 2.0 1.0 5.0 0.5c 1.0 5.0 1.0
87 115 0 22 30 82 22 0 0 0
85 98 0 16 26 74 17 0 0 0
6 DMA N-oxidase activities were calculated from the rate of DMA N-oxide formation at pH 8.4. The composition of the basic reaction media was as described under Methods. The concentration of DMA was 2.5 mu. b va = DMA N-oxidase activity in the presence of the primary amine. v0 = DMA N-oxidase activity in the absence of primary amine. c This is the highest concentration soluble in the reaction medium.
Compound Carbon monoxide” Nitrogenb SKF-525A Cyanide Azide 1-(1-Naphthyl)-2thiourea Catalase Erythrocuprein
IV COMPOUNDS
ACTIVITY
ON THE OF THE
ENZYMES
Concentration Icol/[ozl = 4 in gas phase 100% in gas phase 0.5 rnM 1.0 rnM 1.0 rnM 0.05 rnM 0.50 rnM 1 mg/ml 0.1 mg/ml
$?~~mn 0.0 99 0.0 0.0 0.0
33 92 0.0 0.0
mThe assays were carried out as described in Methods. The concentration of DMA was 2.5 rnM; the reaction time was 2 min. b The reaction vessels were gassed during the 5-min temperature equilibration period. Enzyme, followed by DMA, was added to start the reaction; and the vessels were sealed during the reaction. DMA added prior to temperature equilibration is removed almost quantitatively by a gas flow sufficient to equilibrate the reaction medium.
tested that either have no effect or that inhibit the DMA N-oxidase at high concentrations are octanol, cyclohexanol, nonyl as shown by the information in Table III. As observed with the purified oxidase, n- sulfate, and hexanoic and octanoic acids. Inhibitors. As indicated by the data in octylamine increases the rate of DMA N-oxidation catalyzed by the hepatic Table IV, the amine oxidase is insensitive to microsomes. The increase in pig liver carbon monoxide and SKF-525A, two commicrosomal DMA N-oxidase produced by pounds frequently used to inhibit micro2 mM n-octylamine is usually 2-fold, but somal mixed-function oxidases. The amine 1.3- to 3.0-fold activations have been ob- oxidase activity of the enzyme is also comtained with different preparations of pig pletely insensitive to cyanide and azide even liver microsomes. This is in contrast to the at concentrations 10 times greater than purified oxidase where 2 mu octylamine those listed in the table. The residual cataconsistently stimulates the N-oxidation of lase activity always present in the oxidase is, DMA 1.9-fold. however, inhibited almost 100% by 1 mu Other primary alkylamines will also azide or cyanide. Both of these compounds stimulate the DMA N-oxidase activity of inhibit the oxidase-catalyzed peroxidation of both microsomes and the purified oxidase. methanol and ethanol by hydrogen peroxide. Phenethylamine, amphetamine, and mesca- This activity is probably due to contaminaline are not as effective as n-octylamine; but tion of the purified oxidase by catalase and stimulation of the DMA N-oxidase by these does not appear to be a property of the amines is always observed. Neither ca- amine oxidase. Neither catalase nor erythrodaverine (a diamine) nor aniline affects the cuprein, added in large excessto the reaction DMA N-oxidase activity of microsomes or medium, affects the amine oxidase activity of the purified enzyme. Other compounds of the purified enzyme.
124
ZIEGLER
AND
Of all the compounds tested, l-(lnaphthyl)-2-thiourea appears to be the most selective inhibitor of the amine oxidase, even though this compound would be more accurately described as a substrate with a high affinity for the oxidase but with a very low turnover rate. The more detailed studies of Fok (23) in this laboratory suggest that I-(1-naphthyl)-2-thiourea is competitive with the amine substrate with an apparent Ki of 3 X 10m6M. This compound alone will also slightly stimulate the endogenous NADPH oxidase activity of the purified oxidase, and the naphthylthiourea appears to be consumed during the reaction. This latter interpretation is based on the observation that 0.010-0.015 mM naphthylthiourea, preincubated with NADPH and the oxidase, loses its ability to inhibit the oxidation of DMA. If either NADPH or enzyme is omitted during preincubation, the compound retains its ability to inhibit the oxidation of DMA by the oxidase. It would is appear that 1-(1-naphthyl)-2-thiourea enzymically oxidized to products that do not inhibit the oxidase; but oxidation products of this compound have not, as yet, been identified. While the data are not included in Table IV, 1-(1-naphthyl)-2-thiourea also inhibits the N-oxidation of amines catalyzed by pig or rat hepatic microsomes; but this compound has very little effect on the microsomal-catalyzed oxidative N-demethylation of aminopyrine or propoxyphene. Concentrations of 1-(1-naphthyl)-2-thiourea that inhibit the microsomal N-oxidase activity 80% inhibit the demethylation of the latter two substrates less than 10%. While more work on the effects of 1-(1-naphthyl)-2thiourea on microsomal oxidases will be necessary, these studies indicate that this compound may be a useful t,ool for distinguishing between oxidations catalyzed by the amine oxidase or by other microsomal oxidases. DISCUSSION
The purified oxidase has not been fully characterized as to its amine substrate specificity or its physical properties or ultrastructure. An earlier report (14) with a par-
MITCHELL
tially purified preparation demonstrated that, this oxidase catalyzed the N-oxidation of a large number of synthetic drugs and naturally occuring alkaloids. The highly purified oxidase described in this report N-oxidizes all of the compounds listed in the earlier rcport at increased rates proportional to the increase in the purity of the oxidase. The studies of Sakurai, currently in progress in this laboratory, indicate that the isolated oxidase has a molecular weight close t.o 500,000 and is composed of similar, if not identical, flavoprotein subunits. The precise function and organization of the flavoproteins in the active complex remain to be determined. Earlier studies both in vitro and in viva indicated that cyt. P-450 was not a component of the hepatic mixed-function oxidase that catalyzes the N-oxidation of set- and tertamines. Both carbon monoxide and SKF525A, widely used inhibitors of the microsomal-cyt. P-450 oxidase system, do not inhibit the N-oxidation of amines in vitro (6) ; and in rats the latter compound produced an increased urinary excretion of amine oxides when injected simultaneously with simple dimethylarylor dimethylalkylamines (24). Uehleke et al. (25) have reported that hepatic microsomes, irradiated with uv light to inactivate cyt. P-450, still catalyzed the N-oxidation of secondary and tertiary amines, again indicating that cyt. P-450 was not required for the N-oxidation of these amines. These conclusions are supported by the work presented in this paper. The isolation of a mixed-function amine oxidase free from cytochromes demonstrates that, a microsomal electron transport system is not required for the N-oxidation of set- or ted-amines. A recent direct comparison (26) of the mixed-function amine oxidase and NADPH-cyt. c reductase, both isolated from pig liver microsomes, demonstartes unequivocally that the two flavoproteins are separate enzymes with distinct physical and catalyt,ic properties. Since in microsomes NADPH-cyt. c reductase catalyzes the reduction of cyt. P-450, it would appear that hepatic microsomes contain separate enzyme systems catalyzing the N-oxidat,ion and oxidative-N-dealkylation of many N, Ndimethylalkylor N, N-dimethylaryl-amines.
MICROSOMAL
However, not all microsomal-catalyzed N-oxidations can be attributed to the mixedfunction amine oxidase. The purified amine oxidase does not catalyze the N-oxidation of aniline or of p-chloroaniline amines that are known to be N-oxidized by hepatic microsomes (12). The recent report by Uekleke et al. (25) also suggeststhat different microsomal enzymes were responsible for the cat,:Jytic N-oxidations of primary amines an*f tertiary amines. With the few exceptions described in a recent report from this laboratory (27), it would appear that the N-oxidations of primary amines are catalyzed by a microsomal oxidase different from the mixed-function amine oxidase described in this report. Primary alkylamines, however, do interact with the amine oxidase as shown by the data in Figs. 3 and 4, and Table III. While these amines are not oxidized, they increase the DMA oxidase activity of the enzyme; and the activation of this activity by primary amines is also consistently observed with the microsomal-bound enzyme. Activation of this microsomal oxidase by alkylamines appears to be an inherent property of the enzyme and may reflect a mechanism that controls the activity of this oxidase in vivo. ACKNOWLEDGMENTS We are indebted to Dr. Johnston at Sout,hwestern Medical School, University of Texas at Dallas, for the analysis of the lipids associated with the purified amine oxidase. We also are indebted to Dr. Yukihiko Sakurai for his assistance in perfecting the zone electrophoresis step in the isolation of the oxidase. REFERENCES E. W., AND 1. MACHINIST, J. M., DEHNER, ZIEGLER, D. M. (1968) Arch. &o&m. Biophys. 126, 858. 2. BAKER, J. R., STRUEMPLER, A., AND CHAYKIN, S. (1963) Biochim. Biophys. Acta 71,58. 3. BAKER, J., AND CHAYKIN, S. (1962) J. Biol. Chem. 937, 1309.
OXIDASE
IV
125
4. MASON, H. S. (1957) Science 126, 1185. 5. CONNEY, A. H. (1967) Pharmacol. Rev. 19,317. 6. ZIEGLER, D. M., AND PETTIT, F. H. (1966) Biochemistry 6, 2932. 7. HOPPE-SEYLER, F. A. (1934) Ber. Gesamte Physiol. Exp. Pharmakol. 81, 392. 8. LINTZEL, W. (1934) Biochem. Z. 273,243. R., PHILLIPS, A., TSAI, I., 9. KUNTZMAN, KLUTCH, A., AND BURNS, J. J. (1967) J. Pharmacol. Exp. Ther. 166, 337. 10. BICKEL, M. H. (1969) Pharmacol. Rev. 21, 325. 11. BECKETT, A. H., AND HEWICK, D. S. (1967) J. Pharm. Pharmacol. 19, 134. 12. KIESE, M. (1966) Pharmacol. Rev. 18, 1091. 13. HLAVICA, P., AND KIESE, M. (1969) B&hem. Pharmacol. 18, 1501. 14. ZIEGLER, D. M., MITCHELL, C. H., AND JOLLOW, D. (1969) in Microsomes and Drug
15. 16. 17. 18.
Oxidations (J. R. Gillette, A. H. Conney, G. J. Cosmides, R. W. Estabrook, J. R. Fouts, and G. J. Mannering, eds.), p. 173. Academic Press, New York. DAS, M. L., AND ZIEGLER, D. M. (1970) Arch. Biochem. Biophys. 140, 300. ZIEGLER, D. M., AND PETTIT, F. H. (1964) Biochem. Biophys. Res. Commun. 16, 188. FOK, A. K., AND ZIEGLER, D. M. (1970) Biothem. Biophys. Res. Commun. 41, 534. BROWN, J., AND JOHNSTON, J. M. (1962) J. Lipid
Res. 3, 480. J., AND PETERSON, N. F. (1965) J. Lipid Res. 7, 455. HAUSCHILD-ROGAT, P., AND SMITH, I. (1968) in Chromatographic and Electrophoretic
19. PARKER, 20.
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26.
27.
Techniques. (Ivor Smith, ed.), 2nd edition, p. 475. Wiley (Interscience), New York. DAVIS, B. J. (1964) Ann. N. Y. Acad. Sci. 121, 321. MITCHELL, C. H., AND ZIEGLER, D. M. (1969) Anal. Biochem. 28, 261. FOK, A. K. (1971) Ph.D. Thesis, Department of Chemistry, University of Texas. DEHNER, E. W., MACHINIST, J. M., AND ZIEGLER, D. M. (1968) Life Sci. 7, 1135. UEHLEKE, H., SCHN~TGER, F., AND HELLMER, K. H. (1970) Hoppe Seyler Z. Physiol. Chem. 361, 1475. MASTERS, B. S. S., AND ZIEGLER, D. M. (1971) Arch. Biochem. Biophys. 146, 368. ZIEGLER, D. M., POULSEN, L. L., AND MCKEE, E. M. (1971) Xenobiotica 1,4.