cloning, expression, and regulation of lithocholic acid g

THE JOURNAL OF BIOLOGICAL CHEMISTRY 0 1991 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 266, No. 31, Issue of November 5, pp. 21030-21036,1991 Printed in U. S A .

Cloning, Expression, and Regulation of Lithocholic Acid G@-Hydroxylase*

(Received for publication, January 17, 1991)

Jose TeixeiraS and Gregorio Gilg From the Department of Biochemistry and Molecular Biology, University of Massachusetts Medical School, Worcester, Massachusetts 01655

We have isolated a hamster liver cDNA whose expression is induced upon feeding hamsters with a cholic acid-rich diet. It was identified as a cytochrome P460 family 3 protein, by sequence homology, and named CYP3AlO. The activity of CYP3AlO was determined by transient expression of its cDNA in transfected COS cells and was found to hydroxylate lithocholic acid at position 68. CYP3A10 RNA is 50-fold higher in males than in female hamsters. In males, it appears to be regulated by age with expression highest after puberty. Shortly after weaning (28 days), cholic acid feeding of male hamsters elevates the level of message over that of hamsters fed with normal laboratory chow. Females do not exhibit regulation by cholic acid. In hamster liver, murideoxycholic acid, the 68-metabolite of lithocholic acid, is the major hydroxylated product of lithocholic acid. Lithocholic acid 68- hydroxylase (68-hydroxylase) activity is greatly di- minished in hamster female liver microsomes as would be expected due to the lack of CYP3A10 mRNA in females. Additionally, male liver microsomal 6B-h~- droxylase activity was increased by cholic acid feeding, consistent with the cholic acid-mediated induction of its RNA. These results indicate that, in male hamsters, 68-hydroxylation is the major pathway for detoxification of lithocholate and that, likely, CYP3A10 is responsible for that activity.

The cytochrome P450 (P450)’ superfamily is a group of

* This research was supported in part by American Heart Associ- ation Massachusetts Affiliate Grant-in-aid 13-501-890 and American Heart Association Grant-in-aid 91010730. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) M73992.

$ Predoctoral fellow of the March of Dimes Research Foundation. § Established Investigator of the American Heart Association. To

whom correspondence should be addressed Dept. of Biochemistry and Molecular Biology, University of Massachusetts Medical School, 55 Lake Ave. North, Worcester, MA 01655. Tel.: 508-856-6138 Fax:

The abbreviations and trivial names used are: P450, cytochrome P450; P450 reductase, NADPH-dependent cytochrome P450 reductase; 6a-hydroxylase, lithocholic acid 6a-hydroxylase; 6P-hydroxylase, lithocholic acid 60-hydroxylase; kb, kilobaseb); bp, base pair; HEPES, N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid; CMV, cytomegalovirus; HPLC, high performance liquid chromatography; lithocholic acid, 3a-hydroxy-5P-cholanoic acid; cholic acid, 3a,7a,l2a-trihydroxy-5~-cholanoic acid; chenodeoxycholic acid, 3a, 7a-dihydroxy-50-cholanoic acid; murideoxycholic acid, 3a,6@-dihydroxy-5P-cholanoic acid; hyodeoxycholic acid, 3a,6a-dihydroxy-5@- cholanoic acid; deoxycholic acid, 3a,l2a-dihydroxy-5~-cholanoic acid.

508-856-6231.

hemeproteins that catalyze the oxidation of a variety of en- dogenous and exogenous substrates in concert with NADPH and NADPH-dependent cytochrome P450 reductase (P450 reductase) (Gonzalez, 1989; Nebert et al., 1991). Many P450s are involved in very different metabolic pathways such as steroid hormone metabolism (Gonzalez, 1989; Nebert et al., 1991), drug detoxification (Guengerich, 1987; Shaw et al., 1989; Shimada and Guengerich, 1985), and bile acid synthesis (Bjorkhem, 1985; Bjorkhem and Danielsson, 1974) and are regulated by poorly understood molecular mechanisms that induce or suppress expression or activity. Some P450s are subject to complex endocrine regulation that renders their expression tissue- and sex-specific and/or developmentally programmed (Gonzalez, 1989). The molecular details involved in any of these regulatory mechanisms have yet to be identified.

Among the systems of special interest is the involvement of P450s in cholesterol homeostasis, specifically in converting cholesterol to bile acids for their subsequent elimination from the body (see Fig. 1). Since mammals do not possess enzymes that degrade cholesterol absorbed from the diet, it must be eliminated via the bile acid metabolic pathway (Dietschy, 1984). It is known that 80-90% of intravenously injected cholesterol is catabolized to bile acids (Bjorkhem, 1985). Ath- erosclerosis, gallstone disease, and some lipid storage diseases may be affected by the rate of cholesterol elimination via bile acid biosynthesis and excretion (Bjorkhem, 1985).

Cholic acid and chenodeoxycholic acid are the primary bile acids synthesized by a number of hydroxylations of the steroid nucleus and side chain of the cholesterol molecule (see Fig. 1). These hydroxylations are catalyzed by different liver cytochrome P450s. Primary bile acids are then converted in the gut lumen by microbial enzymes to secondary bile acids such as lithocholic acid and deoxycholic acid by reduction of the 7a-hydroxyl group (Danielsson and Sjovall, 1975).

Lithocholic acid is a toxic bile acid that has been shown to cause cholestasis in experimental animals (Miyai et al., 1975). Indeed, a clinical hallmark of cholestasis is the dramatic increase in serum bile acids, including lithocholic acid. The deleterious effects of this accumulation of lithocholic acid, as well as others, can be attenuated by two detoxification pathways found in the liver: oxidative reactions, mainly hydroxylations, and conjugation with sulfuric or glucuronic acids (Summerfield et al., 1976). Hydroxylation of bile acids makes them more hydrophilic and thus easily excreted in urine or feces. Lithocholic acid can be hydroxylated at position 6 (Fig. I), either below the plane (a) or above the plane (PI, by the enzymes lithocholic acid 6a-hydroxylase (6a-hydroxylase) and lithocholic acid 6P-hydroxylase (6P-hydroxylase) (Hof- man, 1988). In rats, the major hydroxylated product of lithocholic acid is the 6P-metabolite, murideoxycholic acid (Zim- niak et al., 1989).

21030

Lithocholic Acid 6P-Hydroxylase 21031

7a-hydro~ylase Cholesterol

cholcstcrol _____3 7a-hydroxycholesrcrol

Cholic Acid

6p-Hydro~ylasc Cholic Acid .1

3a.6~7a,12a-tctrahydroxy-5~cholan-ZCoic acid

Chenodeoxycholic Acid

I Microbial

dehydroxylases

Lithocholic Acid

Lirhocholic Acid Lirhocholic Acid

6p-Hydroxylase 6a-Hydroxylase / \ Muridcoxycholic Acid Hyodmxycholic Acid

FIG. 1. Bile acid metabolic pathway. A highly schematic de- piction of the bile acid metabolic pathway is shown. Conversion of cholesterol to 7a-hydroxycholesterol by cholesterol 7a-monooxygen- ase (EC 1.14.13.17) is the initial and rate-limiting step in bile acid synthesis. 7a-Hydroxycholesterol undergoes subsequent enzymatic conversions, that have been omitted, in its route to becoming the primary bile acids, cholic acid and chenodeoxycholic acid. Secondary bile acids, lithocholic acid and deoxycholic acid, are synthesized in the gut lumen from the primary bile acids by microbial enzymes and are efficiently reabsorbed into the enterohepatic circulation. In the liver these bile acids can be further hydroxylated for facilitated excretion. Shown are the enzymes and known metabolites involved in the 6-hydroxylation of secondary bile acids.

The perturbation of bile acid metabolism can be a conse- quence of biliary obstruction and alcoholism (Bremmelgaard and Sjovall, 1980; Summerfield et al., 1976), pregnancy (Tho- massen, 1979), fetal development (Strandvik and Wikstrom, 1982), and liver disease and unknown genetic factors (Shoda et al., 1990), whereby the levels of 6-hydroxylated products of bile acids are dramatically elevated in serum and urine. The general evidence for this observation is in clinical cholestasis where the 6-hydroxylation of different bile acids leads to their 6a- and 6P-derivatives. These bile acids are found in substan- tial amounts in the urine and serum of patients with cholestasis and are not found in normal healthy individuals. It would seem logical that 6-hydroxylation can act as a bile acid- induced mechanism for excretion of excess bile acids due to biliary obstruction and liver dysfunction.

In order to better understand how the synthesis and catab- olism of bile acids are regulated and, thus, develop a means for studying bile acid metabolism and the perturbation thereof, we are undertaking an effort to study the regulation of different enzymes involved in bile acid metabolism. We report here the isolation and sequence of a cDNA that encodes lithocholic acid 6P-hydroxylase. In accordance with the recommended nomenclature for P450s (Nebert et al., 1991), this gene has been named CYP3AlO. Expression of this gene is 50-fold higher in males than in females. In young males, expression of CYP3AlO is induced by cholic acid feeding, in contrast to the bile acid-mediated suppression of the rate- limiting enzyme of bile acid synthesis, cholesterol 'Ia-mon- ooxygenase (EC 1.14.13.17) (Jelinek and Russell, 1990; Li et al., 1990; Sundseth and Waxman, 1990).

EXPERIMENTAL PROCEDURES

Material~-~'P-Labeled nucleotides were purchased from Du Pont- New England Nuclear and ["C]lithocholic acid from Amersham Corp. Thin layer chromatography plates were from Merck. Reagents used in cDNA cloning and sequencing were from New England Biolabs, Boehringer Mannheim, U. S. Biochemical Corp., or Gibco/BRL.

Subtractive hybridizations were done with reagents from Invitrogen, San Diego, CA. Authentic murideoxycholic acid was purchased from Steraloids, Wilton, NH and other bile acids from Sigma. Common laboratory chemicals were from Fisher, Sigma, or Bio-Rad. Golden Syrian hamsters were purchased from Charles River Laboratories, Wilmington, MA. DEAE-dextran was from Pharmacia LKB Biotech- nology Inc. Colestipol was purchased from The Upjohn Co.

General Experimental Procedures-Standard recombinant DNA procedures were carried out essentially as described by Sambrook et al. (1989). DNA probes were labeled by random hexanucleotide priming (Feinberg and Vogelstein, 1983) or by primer extension with M13 clones and the universal primer (Church and Gilbert, 1984). Nucleo- tide sequencing was done by the dideoxy chain termination method (Sanger et al., 1977) with M13 universal primers and the Klenow fragment of Escherichia coli polymerase I after subcloning into bac- teriophage M13 vectors (Sambrook et al., 1989). Protein was quantified by the method of Bradford (Bradford, 1976). Total cytochrome P450s were quantified according to Anderson (1985).

Preparation of RNA and Microsomes from Liver-Golden Syrian hamsters were maintained on a 12-h light/l2-h dark cycle, fed either regular laboratory chow or chow supplemented with 0.5% (w/w) cholic acid or 4% (w/w) colestipol for 5 days and were put to death at the middle of the dark cycle. Their livers were removed and used to prepare total cellular RNA by guanadinium thiocyanate homogenization followed by centrifugation through a cesium chloride cushion (Chirgwin et al., 1979). Liver microsomes were prepared by differential centrifugation from the livers of 4-week-old hamsters treated with diets as for RNA preparation. The liver was homogenized in 5 volumes/g homogenization buffer (10 mM Tris acetate, pH 7.4, 1 mM EDTA, and 0.25 M sucrose) with 10 strokes of a Dounce homogenizer. The homogenate was centrifuged at 27,000 X g for 20 min at 4 'C. The supernatant was centrifuged at 100,000 X g for 1 h at 4 "C in a refrigerated ultracentrifuge. The resultant pellet was resuspended in one-fifth the original volume of homogenization buffer and quickly frozen for later use.

Northern and Slot Blot Analyses-Northern blot hybridizations were performed essentially as described (Sambrook et al., 1989). Briefly, 10-pg aliquots of RNA were denatured at 65 "C with glyoxal and dimethyl sulfoxide and electrophoresed overnight through an agarose gel buffered with circulating 10 mM sodium phosphate, pH 7, and blotted onto a nylon membrane overnight. When slot blot analyses were done, 10 pg of RNA was denatured in an equal volume of 18% formaldehyde, 10 X SSC (1 X SSC, 150 mM NaCl, 15 mM sodium citrate) at 65 "C and spotted onto nylon. Blots were dried in a vacuum oven at 80 "C and prehybridized in a 50% formamide hybridization solution at 42 "C. 32P-Labeled probes were added to the solution at 2 X lo6 cpm/ml and hybridized overnight. The blots were washed at 65 "C with 0.1 X SSC plus 0.1% sodium dodecyl sulfate and exposed to x-ray film at -70 "C with intensifying screens. A PetagenTM blot analyzer was used to directly measure counts/min/band or slot.

Preparation, Subtraction, and Screening of Libraries-Poly(A)' RNA (10 pg) was selected from total RNA by two cycles of oligo (dT)-cellulose chromatography (Chirgwin et al., 1979) and used to prepare cDNA libraries from the livers of male hamster fed colestipol (library A) or cholic acid (library B) as described above. After second strand synthesis, BstXI non-palindromic linkers were added by liga- tion, excess linkers removed, and cDNAs ranging in size from 1 to 8 kb size-selected by agarose gel electrophoresis. The selected cDNAs were eluted and ligated to pcDNAJI, a plasmid vector that contains the M13 origin of replication to facilitate single strand synthesis (Duguid et al., 1988). The libraries were transformed into DHla' cells by electroporation generating 2 X lo6 clones each before amplifica- tion.

Libraries were subtracted according to the method of Duguid et al. (1988). Briefly, single-stranded DNA from the library B was biotin- ylated with photobiotin and UV irradiation and hybridized for 24 h with the single-stranded DNA from library A at 68 "C. After hybridization was complete, clones common to both libraries were precipi- tated by addition of streptavidin. The subtracted library was transformed into DHla' cells by electroporation (Potter et al., 1984) and used for screening.

The subtracted library was plated onto nitrocellulose, and replicas of each plate were made. Cells were lysed and the DNA denatured and immobilized on the filter. Filters were prehybridized overnight in 50% formamide at 42 "C according to Sambrook et al. (1989). One set of replicas was probed with hexamer-primed, 32P-labeled cDNA library A subtracted with cholic acid-fed hamster liver mRNA. The other set was probed with cDNA library B subtracted with colestipol-

21032 Lithocholic Acid 6P-Hydroxylase

fed hamster liver mRNA and labeled as above with equal specific activity. After hybridization, filters were washed in 0.1 X SSC, 0.1% sodium dodecyl sulfate at 65 “C and exposed to x-ray film, Plasmids were prepared from putative. positives displaying different signal intensities. These were labeled by random hexanucleotide priming and used as a probe for RNA blots prepared with RNA from colestipol-fed and cholic acid-fed animals. Screening was completed when one clone, pFR29, displayed approximately 4-fold induction with RNA from cholic acid-fed animals versus RNA from normal or colestipol-fed animals.

Subcloning, Sequencing, and Analysis of FR29-pFR29-1 was digested at sites astride the BstXI cloning site of pcDNAII with BamHI and XhoI and the 1.1-kb fragment inserted into M13 vectors. After Sanger dideoxy sequencing (Sanger et al., 1977), a 5’-end 200-bp fragment was isolated by digestion with BstNI and HinfI and used as a probe to rescreen library B to obtain the full-length cDNA. The complete cDNA, pFR29-2, was subcloned by partial digestion with EcoRI, and the 2-kb fragment inserted into the EcoRI site of pCMV (pFR29-5), an expression vector that contains the cytomegalovirus (CMV) promoter and facilitates expression of the cDNA in mammalian cells (Andersson et al., 1989). Complete sequencing of both strands of the full-length cDNA was accomplished by subcloning several fragments into M13 for Sanger dideoxy sequencing. DNA analyses were performed with MicrogenieTM software from Beckman Instruments.

Transient Expression in COS Cells-COS cells were transfected by the DEAE-dextran method (Sambrook et al., 1989). The essentials or modifications are as follows. COS cells were seeded on day 0 at a density of lo6 cells/lOO-mm plate in Dulbecco’s modified Eagle’s medium, 10% fetal calf serum, and 20 mM HEPES. The following day, cells were transfected for 30 min with 10 pg of pFR29-5, pCMV, and/or NADPH-cytochrome P450 reductase cDNA, inserted in the same pCMV vector in 500 pg/ml DEAE-dextran. Cells were then incubated at 37 “C for 3 h in media supplemented with 100 mM chloroquine followed by a 4-min shock with 20% (v/v) glycerol media. After 48 h of incubation at 37 ”C, the cells were washed in ice-cold phosphate-buffered saline and harvested. Microsomes were prepared from the collected cells as for liver except that the first centrifugation was done at 2,000 X g.

Assay of Lithocholic Acid 6-Hydroxylase Activity-Lithocholic acid was prepared for substrate delivery by diluting 7 nmol of 14 Ci/mol [carboxyl-14C]litho~holi~ acid in 20 pl of 1 mg of dilauroyl phospha- tidylcholine/ml of toluene, dried down under NP, and resuspended in 250 mM potassium phosphate, pH 7. Incubations were done under linear conditions in 250 mM potassium phosphate, pH 7 at 37 “C, with 1 mM NADPH and microsomes from COS cells for 1 h or liver microsomes for 20 min. Reactions were stopped by dilution with 500 pl of ethanol, acidified with 20 &l of 0.1 N HCl, and extracted with 5 ml of ethyl ether (Danielsson, 1973). The ether phase was transferred, dried down under Nz, and resuspended in ethanol for spotting on Silica Gel G TLC plates or for reversed phase high performance liquid chromatography (HPLC). The plates were chromatographed with authentic standards in a sealed tank with isooctane:ethyl acetate:acetic acid (5:5:1) (Eneroth, 1963), dried, and exposed to x-ray film overnight. Radioactive spots were excised from the plate and counted in a liquid scintillation counter or else quantified by a Betagen blot analyzer equipped to count I4C. HPLC was done essentially as described by Zimniak et al. (1989). Briefly, the samples were chromatographed on a c18 column with 5 mM potassium phosphate, pH 5, and a 60-min 50-85% methanol gradient and collected. Detection of the murideoxycholic acid standard was done at 214 nm with a Waters 441 detector, and the radioactive reaction products were detected by scintillation counting.

RESULTS

Using differential hybridization techniques (see “Experi- mental Procedures”) we have isolated a hamster liver cDNA (initially called pFR29) whose expression is induced by cholic acid feeding. Fig. 2 is a schematic representation of the isolated cDNAs and the strategy used to determine its nucleotide sequence. The complete nucleotide sequence of pFR29-2 and the predicted amino acid sequence of the protein are shown in Fig. 3. pFR29 encodes a 503-amino acid protein with 76 nucleotides in the 5”untranslated region and 430 nucleotides in the 3”untranslated region. Its deduced amino

0 0.5 1.0 1.5 *’? Kilobascs

PvUU

5’ 3’

pFR29-1 pFR29-2

.c“-( “rv 7 - FIG. 2. Restriction endonuclease map and sequencing strat-

egy for FR29. The scak at the top designates the nucleotide posi- tions (in kilobases) relative to the first nucleotide of the full-length cDNA. The thick black line represents the 503-amino acid coding region, and the 3’- and 5”untranslated regions are represented by a thin line. The restriction endonuclease sites used for subcloning into M13 are indicated. The regions encompassing the initial cDNA isolated (pFR29-1) and the full-length cDNA (pFR29-2) are indicated below the restriction map. The arrows indicate the direction and extent of a given sequencing experiment. Both strands of the entire cDNA were sequenced by the dideoxy chain termination method.

acid sequence contains a putative heme-binding domain pres- ent in all mammalian P450s characterized to date (Gonzalez, 1989) as well as the residues that direct the specificity of P450s for steroids (Gotoh and Fujii-Kuriyama, 1989).

Computer analysis of the sequence revealed that the deduced amino acid sequence of pFR29 was highly homologous to P450 family 3 proteins. This homology approaches 70% with other members of the family previously isolated from human, rat, and rabbit. In Fig. 4, the alignment of the amino acid sequence of FR29 is shown with that of CYP3Al/pcnl, the first of the P450 family 3 proteins whose cDNA was isolated (Gonzalez et al., 1985). Because of this homology, the isolated cDNA has been named CYP3A10 according to the recommended nomenclature system (Nebert et al., 1991).

RNA blot analysis was performed in order to determine the extent of regulation by cholic acid of the steady state level of CYP3A10 mRNA. The autoradiogram of a typical experiment is shown in Fig. 5. The size of the message, 2.1 kb, corresponds to that predicted from the sequence. Regulation of CYP3A10 was determined by measuring the counts/min/band with a Petascope analyzer to ensure accuracy and normalizing to the hybridizable actin message quantified in the same manner (Fig. 5B). Expression of CYP3A10 mRNA is much higher (50-fold) in males (lanes 1-8) than in females (lanes 9-16); in fact, only after exposure of the blot for 4 days (versus 2 h for the males) can CYP3AlO mRNA be detected in female RNA (data not shown). In young males (28 days old) CYP3A10 mRNA was induced approximately 3-fold by cholic acid feeding (lanes 7 and 8 ) in this particular experiment (see other experiments below). There was a slight induction (20%) by cholic acid in males at 7 weeks of age (lanes 5 and 6 ) and no regulation at other ages (lanes 1-4). In females, in addition to the much lower expression of CYP3A10, its RNA is not regulated by cholic acid feeding at any age.

Since family 3 proteins are highly homologous, at least in rat, we eliminated the possibility that our probes might be hybridizing to more than one RNA species under the conditions we normally do the RNA blot analyses (50% formamide hybridization solution and 0.1 X SSC, 65 “C washes) by per- forming a Southern analysis (Fig. 6) with hamster genomic DNA. When we probed one blot with the 5‘-end EcoRI fragment (bp -71 to 186) of the CYP3A10 cDNA (panel A) , two bands were visualized. This was expected since we have

FIG. 3. Nucleotide sequence of the cDNA corresponding to the hamster lithocholic acid GB-hydroxylase mRNA and predicted amino acid sequence of the protein. The nucleotides are numbered on the right-hand side; nucleotide 1 has been arbitrarily defined as the A of the initiator codon. The 5'- untranslated region is numbered nega- tively, and the nucleotides are in lower case letters as are those of the 3'-untranslated region. The last nucleotide is the first A of the poly(A) tail. The amino acid sequence is shown below the nucleotide sequence and numbered with the initiator methionine as the first residue. The amino acid residues of the putative heme-binding domain, characteristic of and highly homologous in all higher eu- karyotic P450s characterized to date (Gonzalez, 1989), are underlined. The residues indicated with an asterisk are those that have been implicated in di- recting the specificity of cytochrome P450s for steroid substrates (Gotoh and Fujii-Kuriyama, 1989).

Lithocholic Acid 6@-Hydroxylase 21033

-71 caggggagccc tgcaagactctcagttgggaggqaaacttg gagccagacctgctgaaggctgaagaaaag -1

ATGGAGCTGATCCCCMCCTTTCTATAGM ACCTGGGTGCTGCTGGCTATCAGCCTGGTG CTCATCTACATATATGGTACCTATTCTCAT 90 MetGluLeuIleProAsnLeuSerIleGlu ThrTrpValLeuLeuAlaIleSerLeuVal LeuIleTyrIleTyrGlyThrTyrSerHis 1 11 21

GGGACTTTTMGMGCTTGGMTTCCTGGA CCAMACCCCTGCCTTTTTTTGGCACCATT TTAWTATCATGATGGTACATGGAAATTT 180 GlyThrPheLysLysLeuGlyIlePrOGly ProLysProLeuProPhePheGlyThrIle LeuGlyTyrHisAspGlyThrTrplysPhe 31 41 51

G A C G A M C G T G C T A T A T A T W X A A A ATATGGGGGTTTTATGACGGTCGAGTGCCT GTGCTGGCTATTGCAGATCCAGAGATCATC 210 AspGluThrCysTyrLysLysTyrGlyLyJ IleTrPGlyPheTyrAspGlyArgvalPro ValLeuAlaI1eA1aAspP~ffiluIleIle 61 71 81

AAAACAGTGCTGGTGAAAGMTGCTACTCA MCTTCACAAACCGTCGGTCTTTTGGGCCA G T G G G A T T T A T G M T C M T C A C C A T A 360 LysThrValLeuvalLysGluCysTyrSer AsnPheThrAsnArgArgSerPheGlyPro ValGlyPheMetLysLysSerIleThrIle 91 101 111

TCTMGGATGMGMTGGAAGAGACTTCGA ACACTACTGTCTCCAGCTTTCACCAGCGGA AAACTCMGAGATGTTCCCCATCATTGGA 450 SerLysAspGluGluTrpLysA=gLeuA=g ThrLeuLeuSerProAlaPheThrSerGly LysLeuLysGluMetPheProIleIleGly 121 131 141

CAGTATGGAGATACGTTGGTGAAAAACTTG AGACGAGAAGAAGAGAAAGGGMGCCCGTC MTATGAAAGAMTCCTTGGAGCTTATAGC 540 GlnTyrGlyAspThrLeuValLysAsnLeu ArgArgGluGluGluLysGlyLysProval AsnMetLysGluIleLeuG1yAlaTyrSer 151 161 171

ATGGATGTGATCACTGGCACATCATTTGGA GTGMTGTCGATTCCCTCMCMCCCAGAG GATCCCTTTGTGCAGMGGCCAGGMGATT 630 MetAspValIleThrGlyThrSerPheGly ValAsnValAspSerLeuAsnAsnProGlu AspPrOPheValGlnLysAlaArgLysIle 181 191 201

TTAAAGTTTMTTTTTTTGATCCCTTTATT CTCTCCATMTACTGTTTCCATTCCTTACC ACMTATATGACTTGTTMGGTTCTCCATT 720 LeuLysPheAsnPhePheAspProPheIle LeuSerIleIleLeuPheProPheLeuThr ThrIleTyrAspLeuLeuArgPheSerIle 211 221 231

T T T C C M G A C M T C M C A M C T T T T T C A M AAATTTA~MCMCGATGMGAAAAATCGC CTTCATTCAMCCAGMGACCCGMTGGAT 810 PheProArgGlnSerThrAsnPhePheLys LysPheIleThrThrMetLysLysAsnArg LeuHisSerASnGlnLysThrArgMetAsP 241 251 261

TTTTTTCAGCTGATGATGMCACTCAGMT TCCAAAGGCAMGAGTCCCAGAAAGCTCTT TCCGATCTAGAAATWAGCACMGCCATT 900 PhePheGlnLeuMetMetAsnThrGlnAsn SerLysGlyLysGluSerGlnLysAlaLeu SerAspLeuGluHetAlaAlaGlnAlaIle 271 281 291

ATTTTCATTTTCGCTGGTTATGMTCCACA AGCACCTCCATTTGTCTCGTATTGTATGM CTGGCCACCCACCCTGATGTGCAGMGAAA 990 IlePheIlePheA1aGlyTyrGluSerThr SerThrSerIleCysLeuValLeuTyrGlu LeuAlaThrHisProAspValGlnLysLys 301 311 321

CTTCACGATGAGATTGACAGCGCTCTGCCC MTMGGCACCTGTCACCTATGATGTCCTG ATGGGCATGGAGTACCTGGACATGGTGATA 1080 leuHisAspGluIleAspSerAlaLe"P~0 ASnLysAlaPrOValThrTyrAspvalLeu MetGlyMetGluTyrLeuAspnetValIle 331 341 351 * MTGMGGTCTCAGATTGTACCCMTTGCT MCAGGCTAGAGAGGATCTCGMGMGGCT GTAGAMTCMTGGACTGTTCATTCCTAM 1170 AsnGluGlyLeuArgLeuTyrProIleAla ASnArgLeuGluArgIleSerLysLysAla ValGluIleAsnGlyLeuPheIleProLys 361 * * * * 371 381

GCGATTACAGTTATGGTACCMCCTATCCT CTTCATCGGSACCCTGAGTACTGGCCAGAG CCTGAGGAGTTCCGCCCTGAAAGGTTCAGC 1260 GlyIleThrValMetValProThrTyrPro leuHisArgASpPcOGluTyrTrpProGlu ProGluG1uPheArgProG1uArgPheSer 391 401 411

AAAGAGMCMGGGCAGCATTGATCCTTAT GTATACATGCCCTTTGGGMCGGACCCAGG MCTGCATTGGCATGAGGTTTGCCCTCCTA 1350 LysG1uAsnLysGlySerIleAspProTyr ValTyrMetPro~eGlyAsn~Prog AsnCyJIleG~MetArgPhe~aLeULeu 421 131 441

AGCATGAAACTGGCTGTCGTCAGTGTCCTG CAGMCTTCACCCTCCAGACTTGTGAGCAA ACAGAGMCCACCTGAAGTTCGCCAGGCM 1440 SerMetLysLeuAlaValValSerVal& GlnAsnPheThrLeuGlnThrCysGluGln ThrGluAsnHisLeuLysPheAlaArqGln 451 461 471

ATMTTCTCCAGCCAGAAAACCCCATCATC CTGMGATCATATCMGAGATAAACCCATA ACCGGAGCATGAatttccctcaggaattat 1530 IleIleLeuGlnProGluA~nProIleIle LeuLysIleIleSerArgAspLysP=oIle ThrGlYAla*** 481 491 501

gctatattcttcaatgggactgtgtcacag aacaccagcaaatt taatt tagcacagaat gtagcacagatgaaattggcttactgaact 1620

ttcattgatgattgagaactcagtacttca ctgaatgtgttcagtgtctatagggagtat tatctattgtgtaattcaatggaaataatt 1710

aagtcaatgatagttgtggagattcatctt ctgattcrcttgtaaccatctacactaatt gcaattagttccatcaattctgagctccaa 1800

ctgt~gta~ctgtttcttt~t~gttct=tc aataatttttatgaaaatagaaactattgt gttgactgttagtaaaaatatercaaactg 1890

t t tct t tgtaaaatt t tatgdtatattgagga tatadataadtatctccttca 1941

observed an intervening sequence 73 bp (see Fig. 3) into the coding region of pFR29-2 that contains an EcoRI site.* When a parallel blot was probed with the 3'-end ScaI fragment (bp 1718 to 2010) of the FR29 cDNA (panel B ) , we observed a single band in each lane. This probe was generated from the 3"untranslated region of the cDNA where one would not expect there to be an intron. This strongly suggests that the conditions we used to do the RNA blot analyses were such that only the product of one gene was being detected however, considering the highly homologous nature of P450s in other species we cannot unequivocally rule out the possibility that

J. Teixeira and G. Gil, unpublished results.

we may be hybridizing to more than one RNA. Additionally, when the same analysis was done under lower stringency conditions (35% formamide hybridization solution and 2 x SSC, 55 "C washes), an innumerable amount of bands were visualized (data not shown), presumably due to hybridization with genes that are homologous to CYP3A10, including other family 3 members.

To determine the nature of the biological activity encoded by CYP3A10, we performed transient expression experiments in COS cells (Fig. 7). We placed CYP3A10 cDNA in the expression vector pCMV (Andersson et al., 1989) that contains the CMV promoter as well as the simian virus 40 (SV40) origin of replication. We transfected COS cells with either the

21034 Lithocholic Acid GB-Hydroxylase FIG. 4. Amino acid alignment of

CYP3AlO/lithocholic acid 68-hydroxylase with that of CYP3A1/ pcnl. Computer analysis of the amino acid sequence encoded by FR29 revealed that it shares a high degree of homology with that of P450 family 3 proteins. Shown is the alignment of lithocholic acid 60-hydroxylase with that of rat pcnl, a well characterized family 3 protein. They share identity (indicated by a I) in 69% of their residues, and only one gap in lithocholic acid 60-hydroxylase was necessary for maximal alignment. The overall homology between rat pcnl and lithocholic acid 60-hydroxylase, considering both identical and conserved (indicated by a :) amino acids increases to 81%.

I

1

76

76

151

151

226

226

300

301

375

376

450

451

HELIPNLSIETnnLAISLVLIYIYGTYSHGTF~LGIPGPKPLPFFGTI~YHDG~DETCY~YGKI~FY

W L L S A L T L E T n n l L Y G F G T R T H G L F K K Q G I P G P K P L P F F G ~ Y ~ L ~ D V E C H ~ Y G K I H G L F I:I: I : : I l I I I I I : I I I : I :I1 :I1 I l l I I I I I I I I I I I I I : I I I I I I I I I I I I I I I I :

D G R W V L A I M P E I I K T C Y S N ~ ~ F G P V G F H l t I G I I :I: I l : I I : I I I I I I I I : I I I I I I I I I I I I I:::::IIIIIII I : I I I I : I l I I : I I I I l I I I ~LFAITDTEHIIMVLVKECFSVFRIRRDFGPVGIffi~VSV~DEE~RY~LSPTFTSGRL~~PIIE

QYGDTLVI(NLRREEEKGKP~EILGAYS~VITGTSFG~SL~PEDPFVQ~KIL~NFFDPFILSI1L

Q Y G D I L V K Y L K Q E ~ T G K ~ ~ G A Y S ~ V I T S T S F G ~ S L ~ P ~ P F ~ K ~ K L L R F D F F D P L F L S ~ I l l1 I l l I: I I I I I I I I : I I I I I I I I I I I I I I I I I I I I I I I I I I 1 : : l : I : l I I I I II::I

FPFLTTIYDLLRFSIFPRQSTNFFKKFITTMKNRLHSNQKTWDFFQmTQN SKGWSQKALSDLEUAAQA

F P F L T P I Y E W L N I ~ P I ( D S 1 E F F ~ ~ T R L D S V O K X R S

I I F I F A G YESTSTSICLVLYELATHPDVQ~LHDEIDS~PNI(AP~YYLD~I~GLRLYP1~LE

IIFIFAGYEPTSSTLSFVLHSY\THPDMIV(1(2EEIDrULPNKAPPTYDRMEHEYLDPNLNETLRLYPIGNRLE

R I S M U V E I N G ~ I P K G I T V H V P T Y P L N R D P E W E P E E F R P E W S I ( E N K G S I D P ~ W F ~ G P ~ C 1 ~ ~

RVCI(I(DVEINGVFWKGSVIPSYUHRDPPllWPEPEEF~ERFSWNKGSIDP~LPFGNGP~C1G~N

LSWKLAWSVLQNFTLQTCEQTENHLI(FM(QI1LQPENPIILKIISRDKPITGA

~LALTKVLQNFSFQPCWTQIPLKLSRQGLLQPTKPIILKVVPRDEIITGS : 1 1 1 1 : I I I I I : I I I I I :I1 : I l l I I I I I : : I I 1 1 1 :

I I I I I II::I :II: I I I I I I : I I II I I I I : I I I I I I I : I I I I l l 1l l l : l : : l l :

I I I I I I I I I 1 1 : : : I I I I I I I I I I I I : I l l I I I I I I I I l l : I I I I I I I I : I I I I I I I I I I I I

I: I I I I I I I : I : I I I I I : I : I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I I : I I I I I I I I I I l I I I I I

A. 1 2 3 4 5 6 7 8 9 10 11 12 13 1 4 1 5 16

kb - 9.4 - 6.6

- 4.4

CW3AIO-

B.

Actin-

- 2.3 - 2.0

I - 2.0 - 2.3

FIG. 5. Regulation of lithocholic acid 68-hydroxylase mRNA. RNA was prepared from the indicated sources and analyzed by Northern analysis as described under “Experimental Procedures.” X/HindI:I-digested DNA was used as a standard and visualized by ethidium bromide staining, and the relative mobilities of the fragments are indicated on the right. Panel A , single-stranded, 3ZP-labeled probes complementary to lithocholic acid 60-hydroxylase mRNA were prepared as described under “Experimental Procedures” from the 5’ EcoRI fragment (bp -71 to 186) of the cDNA and used to detect CYP3AlO RNA. Exposure to x-ray film was done for 1 h at -70 “C with intensifying screens. Panel B, a parallel blot containing the same RNAs was treated as above but hybridized with a single-stranded actin probe (Ma et al., 1986) and exposed for 4 h to film as above.

parental vector pCMV, another construct that contains the P450 reductase cDNA placed in pCMV, our construct containing the CYP3A10 cDNA, or both CYP3A10 and P450 reductase cDNAs. About 48 h after transfection, cells were harvested and microsomes were prepared and assayed for lithocholic acid 6-hydroxylase activity (see “Experimental Procedures”). The reaction products were identified by com- paring their migration distances in a TLC system with those of authentic 6a-hydroxy- (hyodeoxycholic acid) and 68-hy- droxylithocholic acid (murideoxycholic acid). COS cellstrans- fected with vector DNA (lane 1 ) or with P450 reductase cDNA alone (lane 2) produced no activity that converts lithocholic acid. Cells transfected with CYP3A10 cDNA (lane 3) converted 4% of the lithocholic acid used in the assay to a product that migrated with authentic murideoxycholic acid. Cells

CYP3A10

CYP3Al/pcnl

transfected with both CYP3AlO and P450 reductase cDNAs (lane 4 ) converted about 15% of the added substrate to the same product. Two other products were visualized and have yet to be identified. Reversed-phase HPLC confirmed that the major radioactive product of the CYPSA10, reductase- transfected COS cell microsomes assay was murideoxycholic acid with 5% of total counts/min loaded recovered as such (data not shown).

Fig. 7 also shows the lithocholic acid hydroxylase activity of liver microsomes from hamsters of either sex, fed with a control diet or the same diet supplemented with cholic acid (lanes 5-8). Cholic acid feeding, in male hamsters, induced 68-hydroxylase activity by approximately 50-100% (lanes 5 and 6), consistent with the observed induction of CYP3AlO RNA (see Fig. 5). There was no observable induction of 68- hydroxylase activity in females (lanes 7 and 8). Additionally,

A B 5’ end probe 3’ end probe

kb

23 -

” .. .

9.4 - 6.6 -

4.4 -

2.3 - 2.0 -

1 2 3 4 FIG. 6. Southern analysis of hamster genomic DNA with 5’-

end and 3’-end CYP3A10 probes. Hamster genomic DNA was digested with EcoRI (lanes 1 and 3) and XbaI (lanes 2 and 4 ) , electrophoresed through an agarose gel, denatured, neutralized, and transferred to a nylon membrane. The blots were hybridized overnight in 50% formamide solution at 42 “C with single-stranded, 32P-labeled probes (see “Experimental Procedures”) prepared from the 5’ EcoRI, bp -71 to 186 (panel A ) or the 3’ ScaI fragment, bp 1718 to 2010 (panel B ) of FR29 and washed a t 65 “C, 0.1 X SSC. The blots were exposed to x-ray film overnight a t -70 “C with intensifying screens. The relative mobilities of X/HindIII-digested standards are indicated.

Lithocholic Acid GB-Hydroxylase 21035

TRANSFECTED COS CELLS LIVER MICROSOMES

I “ * ‘ “ i W I Reductase

1 2 3 4 5 6 7 8 Lirhocholic

Acid - e

6PHydroxy-

6a-Hydroxy -

origin-

FIG. 7. Lithocholic acid GB-hydroxylase activity in transfected COS cells and in hamster liver microsomes. Lithocholic acid 60-hydroxylase activity was measured as described under “Ex- perimental Procedures” with 200 pg of microsomal protein prepared from COS cells transfected with the indicated DNA or with 20 pg of hamster liver microsomal protein from the indicated sources. The zones corresponding to the migration distance of authentic standards are indicated on the left. The specific activity of 60-hydroxylase was 0.001, 0.001, 0.014, 0.089, 0.339, 0.544, 0.076, and 0.090 nmol/min. mg of protein (lanes 1-8, respectively).

there was an induction of 6a-hydroxylase activity by cholic acid feeding.

A statistical analysis of the cholic acid-mediated induction of CYP3AlO RNA and lithocholic acid GB-hydroxylase activity was done, the results of which are shown in Fig. 8. In this experiment, hamsters (n = 15) were fed either standard laboratory chow or the same diet supplemented with 0.5% cholic acid for 5 days, and their livers were harvested for RNA and microsomes as described under “Experimental Procedures.” Slot blot analyses and GB-hydroxylase assays were done with RNA and microsomes from the individual animals. We observed a 68% increase in CYP3A10 mRNA (solid bars) and a 110% increase in lithocholic acid 6p-hydroxylase activity (stippled bars), p < 0.025 and p < 0.0005, respectively. We also observed a 130% induction of lithocholic acid 6a-hydroxylase (data not shown). The induction in CYP3AlO RNA was less than the %fold observed in Fig. 5, which depicts the results of an experiment with a pool of RNA from three animals. We believe this is due to the high variability of CYP3A10 expression observed in individual animals which necessitated experimentation with a large number of animals and individual analysis to obtain statistically significant data.

DISCUSSION

In the current paper, we report the isolation and nucleotide sequence of a full-length cDNA for the enzyme lithocholic acid GB-hydroxylase. The cDNA has been identified as 6p- hydroxylase based on its ability to program the synthesis of active enzyme upon transfection of the cDNA when placed after the CMV promoter in COS cells (Fig. 7). Other circum- stantial evidence supports this finding. First, by amino acid sequence homology, the isolated cDNA CYP3A10 belongs to family 3 of the P450 gene superfamily (Fig. 4), and all the

0.4 - T T - 0.4 1 I L L

0.3 0.3

0.2 0.2

0.1 0.1

standard 0.5% cholic diet acid diet

FIG. 8. Induction of CYP3A10 mRNA and lithocholic acid GB-hydroxylase activity by cholic acid feeding. RNA and microsomes were prepared from liver as described under “Experimental Procedures” from individual hamsters, 15 of which were fed a standard laboratory diet and 15 fed the same diet supplemented with 0.5% cholic acid. The level of CYP3A10 RNA was quantified by slot blot analysis with a single-stranded probe corresponding to the 5’ EcoRI fragment, bp -71 to 186, in a Petagen (see “Experimental Proce- dures’’) and normalized to the amount of hybridizable actin RNA and shown in arbitrary units (solid bar). Lithocholic acid 60-hydroxylase activity in liver microsomes from the same animals was assayed as described under “Experimental Procedures” with 100 pg of protein and shown in nmol/min.mg (stippled bar). Error bars represent the standard error of the mean. The inductions by cholic acid feeding were statistically significant ( p < 0.025 for RNA; p < 0.0005 for activity).

members of that family characterized to date catalyze hydroxylation of different steroid substrates at position 6 (Gonzalez, 1989). Second, the major hydroxylation product of lithocholic acid in male hamsters is at position 6 (Fig. 7) as it is in both rats (Zimniak et al., 1989) and humans (Radminska et al., 1990). Third, our results in hamster agree with previous reports that cholic acid feeding to rats (Danielsson, 1973) induced lithocholic acid 6p-hydroxylase activity in rat liver microsomes. We have extended those observations by dem- onstrating a concomitant increase in message for CYP3AlO associated with increased lithocholic acid 6p-hydroxylase activity, indicating the induction is at the pretranslational level.

It would not be unexpected for CYP3AlO to have other natural substrates since many P450s have broad substrate specificities, and absolute identification of the preferred or physiological substrate of any P450s may prove a formidable task. However, we have determined the K, for GB-hydroxylase in hamster liver microsomes to be approximately 3 PM (data not shown), within the concentration range reported for lithocholic acid in rat liver (Kurtz et al., 1982). We, therefore, fully expect lithocholic acid to be a physiological substrate for CYP3A10.

Several new insights into bile acid metabolism are deduced from these studies. First, in this study, we unequivocally characterized lithocholic acid GB-hydroxylase as a P450 family 3 protein, CYP3A10. This was done based on the degree of homology with other family 3 proteins (references found in Nebert et al. (1991)). Previous studies hypothesized that 6p- hydroxylation of lithocholic acid could be catalyzed by a P450 family 2 or 3 protein, based on immunological data (Zimniak et dl., 1991). At the amino acid level, CYP3A10 is roughly 70% homologous with the other known members of this family. Due to its activity on lithocholic acid, we believe CYP3A10 is probably a new member of P450 family 3 rather than the hamster homolog of a previously characterized gene.

21036 Lithocholic Acid 6/3-Hydroxylase

The best characterized members of the family, CYP3Al/pcnl and CYP3A5/pcn3, prefer steroid hormone substrates and are not the rat lithocholic acid GB-hydroxylase (Radminska et al., 1990). Absolute identification of CYP3A10 as nonorthologous to other family 3 proteins characterized to date would neces- sitate the isolation of the rat lithocholic acid GB-hydroxylase cDNA.

Second, our results suggest that different genes encode male and female GB-hydroxylase isoenzymes. The level of CYP3AlO/GP-hydroxylase mRNA is 50-fold higher in male uersus female hamsters (Fig. 5), although the level of GB- hydroxylase activity is only &fold lower in females (Fig. 7), which agrees with the 2-fold lower GB-hydroxylase activity in females uersus males described in rats (Zimniak et al., 1991). Interestingly, the level of chenodeoxycholic acid, the imme- diate precursor of lithocholic acid, is significantly higher in female rat liver homogenate (Kurtz et al., 1982; Yousef et al., 1972). This is probably due to the ability of male rat liver to readily convert chenodeoxycholic acid to a more excretable metabolite (Yousef et al., 1973). Sex differences in the enzymes involved in bile acid metabolism have been reported for conjugation (Barnes et al., 1979), oxidoreduction (Bjor- khem, 1985), and bile acid-binding proteins in the liver (LeBlanc and Waxman, 1990), and it certainly would not be surprising if that were the case for GB-hydroxylase. The sex differences we have demonstrated in hamsters, 50-fold higher level of GB-hydroxylase message and 5-fold higher GB-hydroxylase activity in males, are significantly different. The physiological significance of these sex differences in GB-hydroxylase and bile acid metabolism, in general, is unknown.

A third insight derived from these studies relates to the cholic acid-mediated induction of CYP3AlO/G@-hydroxylase mRNA. In young male hamsters, GP-hydroxylase mRNA is induced by cholic acid feeding (Figs. 5 and 8) and no induction thereafter (Fig. 5). This mRNA induction is reflected in the GB-hydroxylase activity (Figs. 7 and 8). Furthermore, cholic acid feeding to hamsters also induces Ga-hydroxylase activity (Fig. 7). Since hydroxylation of lithocholic acid is the major pathway of elimination of this toxic bile acid, the induction of both lithocholic acid hydroxylases suggests that feeding of cholic acid increases the level of lithocholic acid in the hepa- tocytes, which also suggests that a pathway that converts cholic acid into chenodeoxycholic acid may exist. If this is the case one would expect that chenodeoxycholic acid feeding of hamsters also induces GB-hydroxylase, since chenodeoxycholic acid is the direct precursor of lithocholic acid (see Fig. 1). However, preliminary experiments indicate that chenodeoxycholic acid feeding to hamsters suppresses CYP3A10/ GB-hydroxylase RNA, in agreement with a previous report that indicates chenodeoxycholic acid feeding suppresses lithocholic acid GB-hydroxylase activity in rat liver microsomes (Danielsson, 1973). Alternatively, deoxycholic acid (the 7a- dehydroxylated form of cholic acid) may get converted to lithocholic acid by a 12a-dehydroxylation reaction. Experi- ments are in progress to study these possibilities.

Acknowledgments-We wish to thank Carlos B. Hirschberg, Diego Haro, Erica Selva, and Steven L. Smock for discussions, and David J. Waxman for helpful suggestions with the TLC experiments.

REFERENCES Andersson, S., Bostrom, H., Danielsson, H., and Wikvall, K. (1985)

Andersson, S., Davis, D. L., Dahlback, H., Jornvall, H., and Russell,

Barnes, S., Burhol, P. G., Zander, R., Haggstrom, G., Settine, R. L.,

Methods Enzymol. 1 1 1 , 364-377

D. W. (1989) J. Biol. Chem. 264,8222-8229

and Hirschowitz, B. I. (1979) J. Lipid Res. 20,952-959 Bjorkhem, I. (1985) in Sterols and Bile Acids (Davidson, H., and

Sjovall, J., eds) pp. 231-278, Elsevier Scientific Publishing Co., Amsterdam

Bjorkhem, I., and Danielsson, H. (1974) Mol. Cell. Biochem. 4 , 79-95 Bradford, M. M. (1976) Anal. Biochem. 72 , 248-254 Bremmelgaard, A., and Sjovall, J. (1980) J. Lipid Res. 21, 1072-1081 Chirgwin, J. M., Przybyla, A. E., MacDonald, R. J., and Rutter, W.

Church, G. M., and Gilbert, W. (1984) Proc. Natl. Acud. Sci. U. S. A.

Danielsson, H. (1973) Steroids 2 2 , 667-676 Danielsson, H., and Sjovall, J. (1975) Annu. Reu. Biochem. 44 , 233-

Dietschy, J. M. (1984) Klin. Wochenschr. 6 2 , 338-345 Duguid, J. R., Rohwer, R. G., and Seed, B. (1988) Proc. Natl. Acud.

Eneroth, P. (1963) J. Lipid Res. 4 , 11-16 Feinberg, A. P., and Vogelstein, B. (1983) Anal. Biochem. 132,6-13 Gonzalez, F. J. (1989) Pharmacol. Reu. 4 0 , 243-288 Gonzalez, F. J., Nebert, D. W., Hardwick, J. P., and Kasper, C. B.

(1985) J. Biol. Chem. 260,7435-7441 Gotoh, O., and Fujii-Kuriyama, Y. (1989) in Basis and Mechanism of

Regulation of Cytochrome P450 (Ruckpaul, K., and Rein, H., eds) pp. 196-243, Akademie-Verlag, Berlin

Guengerich, F. P. (1987) in Mammalian Cytochrome P-450 (Guenger- ich, F. P., ed) pp. 1-54, CRC Press, Inc., Boca Raton, FL

Hofman, A. F. (1988) in The Liver: Biology and Pathobiology (Arias, I. M., Jakoby, W. B., Popper, H., Schachter, D., and Shafritz, D. A., eds) pp. 553-572, Raven Press, New York

Jelinek, D. F., and Russell, D. W. (1990) Biochemistry 29,7781-7785 Kurtz, W., Leuschner, U., Hellstern, A., and Janka, P. (1982) Hepato-

LeBlanc, G. A., and Waxman, D. J. (1990) J. Biol. Chem. 265,5654-

Li, Y. C., Wang, D. P., and Chiang, J. Y. L. (1990) J. Biol. Chem.

Ma, P. T. S., Gil, G., Siidhof, T. C., Bilheimer, J. L., Goldstein, J. L.,

8374 and Brown, M. S. (1986) Proc. Natl. Acad. Sci. U. S. A. 83, 8370-

Miyai, K., Mayr, W. W., and Richardson, A. L. (1975) Lab. Invest.

Nebert, D. W., Nelson, D. R., Coon, M. J., Estabrook, R. W., Feyer- eisen, R., Fujii-Kuriyama, Y., Gonzalez, F. J., Guengerich, F. P., Gunsalus, I. C., Johnson, E. F., Loper, J. C., Sato, R., Waterman, M. R., and Waxman, D. J. (1991) D N A ( N . Y . ) 10 , l -14

Potter, H., Weir, L., and Leder, P. (1984) Proc. Natl. Acud. Sci.

Radminska, A., Zimniak, P., Falany, J., Iscan, M., Lester, R., and Waxman, D. J. (1990) FASEB J. 4 , A1972 (abstr.)

Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed., Cold Spring Harbor Lab- oratory, Cold Spring Harbor, NY

Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74,5463-5467

Shaw, P. M., Barnes, T. S., Cameron, D., Engeset, J., Melvin, W. T., Omar, G., Petrie, J. C., Rush, W. R., Snyder, C. P., Whiting, P. H., Wolf, C. R., and Burke, M. D. (1989) Biochem. J. 263,653-663

Shimada, T., and Guengerich, F. P. (1985) Mol. Pharmacol. 28,215- 219

Shoda, J., Tanaka, N., Osuga, T., Matsuura, K., and Miyazaki, H. (1990) J. Lipid Res. 31, 249-258

Strandvik, B., and Wikstrom, S.-A. (1982) Eur. J. Clin. Inuest. 12 ,

Summerfield, J. A., Billing, B. H., and Shackleton, C. H. L. (1976)

Sundseth, S. S., and Waxman, D. J. (1990) J. Biol. Chem. 2 6 5 ,

Thomassen, P. A. (1979) Eur. J. Clin. Invest. 9,425-432 Yousef, I. M., Kakis, G., and Fisher, M. M. (1972) Can. J. Biochem.

Yousef, 1. M., Magnusson, R., Price, V. M., and Fisher, M. M. (1973)

Zimniak, P., Holsztynska, E. J., Lerster, R., Waxman, D. J., and

Zimniak, P., Holsztynska, E. J., Radominska, A., Iscan, M., Lester,

J. (1979) Biochemistry 18 , 5294-5299

81,1991-1995

253

Sci. U. S. A. 8 5 , 5738-5742

gastroenterology 29, 227-231

5661

265,12012-12019

32,527-535

U. S. A. 81,7161-7170

301-305

Biochem. J. 154,507-516

15090-15095

50,402-408

Can. J. Physiol. Pharmacol. 5 1 , 418-423

Radominska, A. (1989) J. Lipid Res. 30,907-918

R., and Waxman, D. J. (1991) Biochem. J. 275,105-111

cloning, expression, and regulation of lithocholic acid g

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