A COMPARATIVE STUDY OF XENOBIOTIC-METABOLIZING ENZYMES IN LIVER AND INTESTINE OF VARIOUS ANIMAL SPECIES

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1 DRUG METABOLISM A DIsPOsITIoN Copyright #{174} 1974 by The American Society for Pharmacology and Experimental Therapeutics Vol. 2 No. S Printed in U.S.A. A COMPARATIVE STUDY OF XENOBIOTIC-METABOLIZING ENZYMES IN LIVER A INTESTINE OF VARIOUS ANIMAL SPECIES RAJERA S. CHHABRA. ROBERTA J. POHL. A JAMES R. FOUTS Pharmacology Branch. National Institute of Environmental Health Sciences (Received May ) ABSTRACT s, mice, hamsters, guinea pigs. and rabbits were used in a preliminary study to select a model animal for more detailed study of xenobiotic-metabolizing enzymes in intestine. The xenobiotic oxidation reactions studied included hydroxylation of aniline, biphenyl, and benzpyrene and N-demethylation of ethylmorphine. NADPH-cytochrome c reductase activity and cytochrome P-450 were measured as parts of the microsomal electron transport system. The xenobioticmetabolizing enzymes were present in livers of all the and varied in activity over a 2- to 6-fold range among for any given enzyme. In intestines from mice, rats, guinea pigs. and hamsters, either some of the enzymes were absent or had very low activity which would require very sensitive methods for detection. The rabbit emerged as the best animal for studying intestinal microsomal xenobiotic metabolism since all xenobiotic-metabolizing enzymes studied were present in easily measurable quantities. Recently, there has been a great deal of interest in the study of drug metabolism in extrahepatic tissues such as lung, skin, and intestine (1-4). The study of detoxication-toxication mechanism in cxtrahepatic tissues is important since some of these tissues can be the portal of entry for environmental pollutants and it is important to know about the fate of environmental contaminants or drugs at the first lines of defense in the body. Small intestine is one of the extrahepatic tissues which has been shown to metabolize foreign substances, although the rates of metabolism reported are generally lower than those observed in liver. Studies on mixed-function oxidases in intestine have often been limited to only one mixed-function oxidase activity, i.e. benzpyrene hydroxylase, and this has usually been studied only in rats (5-8). A systematic study of mixed-function oxidases in intestine is needed to better compare these with analogous systems in liver. Furthermore such a study may help to reveal the role of mixed-function oxidases in the toxication-detoxication of a variety Send reprint requests to: Rajendra S. Chhabra. Ph.D.. Pharmacology Branch. National Institute of Environmental Health Sciences. P.O. Box Research Triangle Park. NC of drugs and foreign compounds. The present research was an attempt to find the most suitable animal model for a systematic comparison of mixed-function oxidases of liver and intestine. Experimental Procedure Materials. Five- to seven-week-old male mice of the CDI strain weighing g, and male rats of the CD strain weighing g were obtained from Charles River Breeding Laboratories, Wilmington, Mass. Male Syrian golden hamsters weighing g were purchased from Lakeview Colony, Newfield, N.J. Male New Zealand White rabbits weighing kg were supplied by Arrow Farm, Statesville, NC. Male Hartley guinea pigs were bred at this institute and weighed about g. s, mice, and hamsters were fed with Wayne Lab-Blox and water ad libitum. s and guinea pigs had uncontrolled access to National Institutes of Health Feed A and water. Feed was analyzed at intervals for residual chlorinated pesticides and estrogen levels. No detectable amounts of estrogen were allowed and chlorinated pesticides did not exceed more than 0.03 ppm of feed. All animals were housed in stainless steel hanging cages except mice which were kept five per plastic box with Absorb-Dri bedding. Animals were kept in the animal house for approximately days before death. The animals were killed between 8:30 and 9:30 am. Preparation of Microsomes and Assays. s, hamsters, mice, and guinea pigs were killed by 443

2 444 CHHABRA ET AL. cervical dislocation, whereas rabbits were killed by air embolism. Livers were removed and placed in beakers containing ice-cold KCI-HEPES buffer, ph 7.4 (1.25 mm N-2-hydroxyethylpiperazine-N -2-ethanesulfonic acid in 1.15% KCI). The entire small intestine was removed and washed 2-3 times by forcing ice-cold saline (0.9% NaCI) from a syringe through the lumen until the intestines were visibly free from excreta. The intestines were then cut longitudinally with scissors and laid on the convex surface of a watch glass, and the mucosal layer was scraped out with a spatula. These mucosal scrapings were kept in ice-cold beakers until homogenization. Small intestine mucosa from 4-10 animals were pooled with all except rabbit; no pools were needed when rabbit intestines were used. Liver or intestinal mucosal scrapings were homogenized in 2 parts of KC1-HEPES using a Potter-type homogenizer with a plastic pestle. The microsomal fractions were prepared by methods described previously (9). In experiments related to stabilization of cytochrome P-450 in rat intestinal microsomes, the salt aggregation method of Kamath et a!. for preparing hepatic microsomes was used with a slight modification (10, 11). The intestinal mucosal scrapings were homogenized in 4-5 parts of 0.25 M sucrose in a Potter-type homogenizer. The lo,000g supernatant fraction (obtained from centrifugation of the homogenate) was diluted 5 times with aggregating solution containing 12.5 mm sucrose, 8 mm CaCI,, and 5 mm MgCl2, and the mixture was centrifuged to harvest the microsomes (20 mm at l0,000g in a Sorvall RC-2B refrigerated centrifuge using the GSA rotor). The supernatant fraction was discarded; the pellet was then resuspended in KCI-HEPES buffer (ph 7.4) for enzymic assays. In vitro assay of N-demethylation of ethylmorphine was performed by estimating the formaldehyde formed (12). Aniline hydroxylase was assayed as described previously (9). Bcnzpyrene hydroxylase activity in the microsomal fraction of tissues was determined as described by Hansen and Fouts (13). Biphenyl hydroxylation was measured by the formation of 4-hydroxybiphenyl (14). The typical incubation mixture for all these enzyme reactions contained glucose 6-phosphate and MgSO4 (12.5 zmo1 each); NADP, 2.5 tmol; 0.5 ml of 0.5 M HEPES buffer, ph 7.4; glucose 6-phosphate dehydrogenase, 2 units (Sigma Chemical Co., St. Louis, Mo.); and microsomal protein, 2.5 mg; the total incubation volume was 2.5 ml. The final substrate concentrations for ethylmorphine N-demethylase, benzpyrene hydroxylase, and biphenyl hydroxylase were 10, 0.6, and 60 mm, respectively. The reaction mixtures were incubated for 15 mm in air at 37#{176}C. NADPH-cytochrome c reductase activity was assayed at 37.5#{176}Cby recording the appearance of reduced cytochrome c at 550 nm in a Gilford model 240 spectrophotometer (15); incubation mixtures contained 0.3 M HEPES buffer, ph 8.0, 1.6 mm KCN, 167 mm KC1, 0.08 mm cytochrome c (Sigma type III), 0.6mg of microsomal protein, and 0.3mM NADPH in a total volume of 3 ml. Cytochrome P-450 content of the microsomal preparations suspended in 0.1 M HEPES buffer, ph 7.6, was assayed by the method of Omura and Sato (16) in an Aminco DW-2 spectrophotometer. The molar extinction coefficient of 91,000 M cm was used for estimation of microsomal cytochrome P-450 concentrations. Results Table 1 shows the comparison of ethylmorphine N-demethylase activity in hepatic and intestinal microsomes from. This enzyme activity was found to be present in livers from all with activity varying over a 5-fold range. However, intestinal m icrosomal N-demethylase activity was detected only in rabbit and guinea pig. Biphenyl and benzpyrene hydroxylase activities were present in both hepatic and intestinal microsomes of all studied (tables 2 and 3, respectively). Hepatic benzpyrene hydroxylase activity varied among over a 5-fold range, whereas intestinal benzpyrene hydroxylase showed a 25-fold variation among. Biphenyl hydroxylase showed a 3-fold variation in hepatic and a 4-fold variation in intestinal microsomal activity among. As with ethylmorphine N-demethylase, aniline hydroxylase was found to be present only in rabbit and guinea pig intestinal microsomes (table 4). NADPH-cytochrome c reductase activity was found to be present both in hepatic and intestinal microsomes of all the under investigation. In both liver and intestine, NADPHcytochrome c reductase varied over a 3-fold range among (table 5). Table 6 shows the concentrations of cytochrome P-450 in microsomes from liver and intestine of the under study. Cytochrome P-450 was present in both tissues of all the except for rat intestine where it was TABLE Ethylmorphine N-demethylase activity in liver and intestine of Activity is expressed as nanomoles of formaldehyde formed per mm per mg of microsomal protein ± SE ± 22.8(3) 37.7 ± 3.4(6) ± 8.9 (4) ± 9.6(4) ± 15.1(4) I 11.2 ± 1.6(3) 8.8 ± 0.3(4) 5 #{176}Number of animals or separate tissue pools is given in, activity not detected. The least detectable amount of HCHO is 4 nmol/mg of microsomal protein in 15 mm

3 INTESTINAL DRUG METABOLISM 445 TABLE 2 Biphenyl hydroxylase activity in liver and intestine of Activity is expressed as nanomoles of 4-hydroxybiphenyl formed in 15 mm per mg of microsomal protein ± SE. Guinea pig 58.1 ± 1.6(4) ± 1.8(4) 34.4±1.4(4) 95.3 ± 10.0(4) 47.4 ± 3.0(4) 8.2 ± 0.6(4) 12.8 ± 2.9(4) 3.2 ±0.3(4) 8.6 ± 0.7(4) 3.2 ± 0.5 (4) The least detectable amount of 4-hydroxybiphenyl is 0.5 nmol/mg of microsomal protein in 15 mm. TABLE 3 Benzpyrene hydroxylase activity in liver and intestine of Activity is expressed as relative fluorescence units in is mm per mg of microsomal protein ± SE. A quinine sulfate solution (3.ag/ml in 0.1 NaOH) was used as a fluorescent standard with each experiment ± 191 (4)0 995 ± 37(4) 3009±211(4) 1705 ± 172(4) 579 ± 41(4) 835 ± 39(4) 372 ± 86(4) 138±8(4) 101 ± 21(4) 33 ± 6(4) The least detectable amount of benzpyrene hydroxylase activity is 10 units of relative fluorescence per mg of microsomal protein in IS mm. TABLE 4 A niline hydroxylase activity in liver and intestine of Activity is expressed as nanomoles of p-aminophenol formed in 15 mm per mg of microsomal protein ± SE. Guinea pig 9.8 ± 2.3(3) ± 1.0(6) 15.0±0.9(4) 24.1 ± 1.9(4) 19.1 ± 1.3(4) 2.0 ± 0.2 (3) 2.1 ± 0.7 (4) Number of experiments or separate tissue pools is given in, activity not detected. The least detectable amount of p-aminophenol formed is 0.8 nmol/mg of microsomal protein per IS mm TABLE 5 NA DPH-cytochrome c reductase activity in liver and intestine of Activity is expressed as nanomoles of cytochrome c reduced per mm per mg of microsomal protein ± 15.S (4) ± 2.8(4) 118.3±18.3(4) ±6.8(4) 44.3 ± 4.1 (4) ± 23.0(4) 43.6 ± 4.6(4) 49.7±7.3(4) 90.0± 1.6(4) 26.9 ± 2.6(4) Activity in at of That in Liver The least detectable amount of cytochrome c reduced is 1.0 nmol/mg of protein per mm. TABLE 6 Cytochrome P-450 content in liver and intestine of Content is expressed as nanomoles per mg of microsomal protein ± SE. Guineapig Mice 1.1 ± 0.32(4) 1.4S ±0.16(4) 0.84 ± 0.07 (4) 1.1 ±0.07(4) 1.26 ± 0.07(4) 0.38 ± 0.07 (4) 0.18 ±0.02(4) ±0.0I(4) 0.16 ± 0.01 (4) Amount in Intestineas That in Liver For explanation see Results. The least detectable amount of cytochrome P-450 is 0.01 nmol/mg of microsomal protein. detected in its inactivated form, cytochrome P-420. s had the highest concentrations of intestinal microsomal cytochrome P-450 of all studied Attempts to Stabilize Cytochrome P-450 in Intestinal Microsomes. was the only studied which did not show a detectable concentration of cytochrome P-450 in intestinal microsomes, although a considerable absorption peak in the dithionite difference spectrum occurred at nm (fig. 1,!ine A). This peak was not due to contamination of intestinal microsomes by hemoglobin. Therefore, the peak at nm appeared to derive from conversion of cytochrome P-450 to cytochrome P-420. The preparation of rat intestinal microsomal fractions by the salt aggregation method appeared to prevent some degradation of cytochrome P-450 to cytochrome P-420 (fig. 1,!ine B).

4 446 CHHABRA ET AL. S U IC 450 m,u FIG. 1. Cytochrome P450 in rat intestinal mucosa Suspensions contained 3 mg microsomal protein/mi in 0.1 M HEPES buffer, ph 7.6, in a total volume of 3 ml. Sample cuvettes were bubbled for sec with alkaline dithionite-scrubbed CO. A few milligrams of sodium dithionite were added to both sample and reference cuvettes. Scans were repeated until maximum absorbance at 450 nm was recorded. Curve A, microsomes isolated by conventional method. Curve B, microsomes isolated by salt-aggregation method. Several attempts (listed below) were made, without success, to block the total conversion of cytochrome P-450 to cytochrome P-420 in rat intestinal homogenates during preparation of microsomes. 1) Trypsin might be converting cytochrome P-450 to P-420 during homogenization; therefore, trypsin inhibitor (Sigma type 1-S) was added to the homogenizing solution at I and 5 mg/g of rat intestinal mucosa, and remained throughout the preparation of microsomes. 2) Metals such as Hg2 might catalyze the denaturation of cytochrome P-450. Therefore, EDTA (1 mm) was added to the homogenizing medium and throughout the preparation of microsomes. 3) s are the only we studied that do not have a gallbladder, and bile salts in intestine may be involved in degrading cytochrome P-450. Several experiments were unable to demonstrate this however: a) s starved 36 hr before death (to decrease bile flow) did not have additional (detectable) amounts of cytochrome P-450 when compared to rats with access to food. b) Oral administration of the inhibitor of bile production, 1-naphthyl isothiocyanate, or surgical bile duct ligation did not block the conversion of intestinal cytochrome P-450 to cytochrome P-420. The following experiments showed that a cornponent in rat intestinal microsomes was degrading cytochrome P ) Mixing rat intestinal mucosa with rabbit intestinal mucosa at the homogenization step in the preparation of microsornes caused the loss of 40% of the rabbit intestinal cytochrome P-450, with the appearance of a peak in the dithionite difference spectrum at 420 nm. Similarly mixing rat intestinal microsomes with rabbit intestinal microsomes I V hr before assay of cytochrome P-450 caused a loss of 25% of the rabbit intestine cytochrome P-450 with the appearance of a peak at 420 nm. 2) The time-course of rat intestinal microsomally catalyzed degradation of rat hepatic microsomal cytochrorne P-450 showed a loss of 10% in 50 mm when compared to controls of rat liver microsomes mixed with bovine serum albumin equivalent to the protein concentration of intestinal microsornes. No further attempts were made to identify the component which degrades cytochrome P-450 in rat intestinal microsornes. Discussion Environmental contaminants come in contact with small intestine either through ingestion or inhalation (by swallowing). Wattenberg has studied in detail the metabolism of one of the environmental contaminants (benzpyrene) in rat intestine (5-7). His work has shown that rat intestinal microsomes may play an important role in detoxication of arylhydrocarbons (17). Recent studies have also shown that aryihydrocarbon hydroxylase activ ity in rat intestinal microsomes may depend on exogenous inducers present in the diet (8). There have been very few reports on studies of extrahepatic mixed-function oxidases in other than rats. In the present study, rabbits, guinea pigs, hamsters, rats, and mice were used to study a number of intestinal mixed-function oxidases in order to select a suitable, with detectable amounts of several of these enzymes, for further studies of xenobiotic-metabolizing enzymes in intestine. The results of our present study show that there are qualitative as well as quantitative differences in activities of xenobiotic-metabolizing enzymes in microsomes from the small intestines of. The activities of several mixed-function oxidases were generally lower in intestinal than hepatic microsomes of all the (range from 4

5 INTESTINAL DRUG METABOLISM 447 to 80% of that found in liver). Ethylmorphine N-demethylase and aniline hydroxylase were not detected in intestinal microsomes of rats, mice, and hamsters, although the intestinal microsomal N-demethylation of aminopyrine and 3-methyl-4- methylaminoazobenzene has been reported by other workers (18, 19). The present study also reveals that the rat may not be an ideal animal for the study of a variety of intestinal mixed-function oxidations since many mixed-function oxidase activities were either absent or required very sensitive methods for detection. The absence of most of the mixed-function oxidases in rat intestinal microsomes might have been due to the conversion of most of the cytochrome P-450 to the inactive form (P-420) during homogenization of rat intestine, or to have been due to the presence of a relatively specific cytochrome P-448 system as suggested by Mannering (8). seemed to be a most suitable animal for studies of mixed-function oxidases in intestine. All xenobiotic-metabolizing enzymes studied were present in rabbit intestinal microsomes and were detectable within the limits of the methods employed for enzymic assays. The further characterization of xenobiotic-metabolizing enzymes in rabbit intestinal microsomes is currently under investigation. References 1. J. R. Bend, G. E. R. Hook, R. E. Easterling, T. E. Gram, and J. R. Fouts, J. Pharmacol. Exp. Ther. 183, 206 (1972). 2. F. J. Wiebel, J. C. Leutz, and H. V. Gelboin, Arch. Biochem. Biophys. 154, 292 (1973). 3. K. Hartiala, Pharmacol. Rev. 53, 496 (1973). 4. R. M. Welch, J. Cavallito, and A. Loh, Toxicol. AppI. Pharmacol. 23, 749 (1972). 5. L. W. Wattenberg, J. L. Leong, and P. J. Strand, CancerRes. 22, 1120(1962). 6. L. W. Wattenberg, Cancer 28, 99 (1971). 7. L. W. Wattenberg, Toxicol. AppI. Pharmacol. 23, 741 (1972). 8. N. G. Zampaglione and G. J. Mannering, J. Pharmacol. Exp. Ther. 185, 676 (1973). 9. R. S. Chhabra, T. E. Gram, and J. R. Fouts, Toxicol. App!. Pharmacol. 22, 50 (1972) A. Kamath, P. A. Kummerow, and K. A. Narayan, F.E.B.S. Leti. 17, 90 (1971). 11. S. A. Kamath and E. Rubin, Biochem. Biophys. Res. Commun. 49, 62 (1972). 12. T. Nash, Biochem. J. 55, 416 (1953). 13. A. R. Hansen and J. R. Fouts, Biochem. Pharmacol. 20, 3125 (1971). 14. P. J. Creaven, D. V. Parke, and R. T. Williams, Biochem. J. 96, 879 (1965). 15. M. A. Peters and J. R. Fouts, J. Pharmacol. Expt. Ther. 173, 233 (1970). 16. T. Omura and R. Sato, J. Biol. Chem. 239, 2370 (1964). 17. L. W. Wattenberg, Progr. Exp. Tumor Res. 14, 89 (1970). 18. F. B. Thomas, N. Baba, N. J. Greenberger, and D. Salsbury, J. Lab. Clin. Med. 80, 548 (1972). 19. R. E. Billings and L. W. Wattenberg, Proc. Soc. Exp. Biol. Med. 139, 865 (1972).