THE J O L R N A L OF RlOLOClCAL CHEMISTRY Val. 266, No. 15, Issue of May 25, pp. 9413-9418, 1991 Printed in U, S. A.

Effect of Hydrophobic Bile Acids on 3-Hydroxy-3-Methylglutaryl- Coenzyme A Reductase Activity and mRNA Levels in the Rat*

(Received for publication, January 2, 1991)

Paul F. Duckworth$, Z. Reno VlahcevicS, Elaine J. StuderQ, Emily C. GurleyQ, Douglas M. Heurnan$, Zafarul H. BegTl, and Phillip B. HylemonQII From the Departments of $Medicine, and §Microbiology and Immunology, Medical College of Virginia, Virginia Commonwealth University, Richmond, Virginia 23298 and the TlMolecular Disease Branch, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892

We have previously reported that relatively hydro- phobic bile acids, decreased hepatic 3-hydroxy-3- methylglutaryl-coenzyme A reductase (reductase) ac- tivity whereas, hydrophilic bile acids had little effect on the enzyme. The purpose of the present study was to determine in more detail the mechanism of down- regulation of hepatic reductase activity by hydropho- bic bile salts. Groups of rats were fed bile acids of differing hydrophobicity: ursodeoxycholic, cholic (CA), chenodeoxycholic (CDCA), deoxycholic (DCA), or cholesterol for 14 days. Reductase specific activities and concentrations of reductase protein were deter- mined in hepatic microsomes. Quantitation of “steady state” levels of reductase mRNA was performed using Northern and dot blot hybridization. Reductase gene transcriptional activity (nuclear “run-on”) was deter- mined in nuclei isolated from livers of animals fed different bile acids. Hydrophobic bile acids and choles- terol significantly decreased reductase activity: CA (57%), CDCA (77%), DCA (73%), cholesterol (89%), and reductase protein levels as measured by an en- zyme-linked immunosorbent assay method were also decreased; CA (27%), CDCA (31%), DCA (42%), and cholesterol (35%). Reductase mRNA levels were also decreased after feeding hydrophobic bile acid: CA (43%), CDCA (47%), DCA (54%), and cholesterol (53%). Ursodeoxycholic, a hydrophilic bile acid, caused a much smaller decrease in reductase activity (18%), protein mass (l6%), and mRNA levels (10%).

Decreased transcriptional activities were observed in CA- and cholesterol-fed rats. Surprisingly, CDCA- and DCA-fed animals showed transcriptional activities similar to control animals even though steady state mRNA levels were low in CDCA- and DCA-fed ani- mals. We hypothesize a post-transcriptional regulation of reductase mRNA by hydrophobic bile acids.

The activities of HMG-CoA’ reductase, the rate-limiting

*This work was supported by Grant PO1 DK38030 from the National Institutes of Health. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “aduertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

)I To whom correspondence should be addressed: Dept. of Micro- biology and Immunology, Box 678 MCV Station, Richmond, VA 23298-0678.

’ The abbreviations used are: HMG-CoA, 3-hydroxy-3-methyl- glutaryl-coenzyme A; CA, cholic; CDCA, chenodeoxycholic; DCA, deoxycholic; ELISA, enzyme-linked immunosorhent assay; SDS, sodium dodecyl sulfate; EGTA, [ethylenebis(oxyethylenenitrilo)]tet- raacetic acid; HEPES, 4-(2-hydroxyethyl)-l-piperazineethanesul- fonic acid; TES, 2-([2-hydroxy-l,l-bis(hydroxymethyl)ethyl]amino~- ethanesulfonic acid.

enzyme in the cholesterol biosynthetic pathway, and choles- terol 7a-hydroxylase, the rate-limiting enzyme in the bile acid biosynthetic pathway, often vary in a parallel fashion under a variety of experimental and physiological conditions (1, 2). We have previously reported that feeding of relatively hydro- phobic bile acids (cholic, chenodeoxycholic, and deoxycholic acid) decreased the activities of both cholesterol 7a-dehydrox- ylase and HMG-CoA reductase in roughly similar proportions and in order of increasing hydrophobicity (cholic < cheno- deoxycholic < deoxycholic) (3-6). The activities of these two enzymes were highly correlated with the hydrophobic-hydro- philic balance (hydrophobicity index) of the circulating bile salt pool as measured by reverse-phase high performance chromatography. In contrast, the activity of acyl CoA:cholesterol acyltransferase, another microsomal enzyme involved in cholesterol metabolism, was not correlated with the hydrophobic-hydrophilic balance of the bile salt pool suggesting that the observed effects are specific for HMG- CoA reductase and cholesterol 7a-hydroxylase (6). Hydro- philic bile acids (ursocholic, ursodeoxycholic, hyocholic, and hyodeoxycholic) had no effect on either HMG-CoA reductase or cholesterol 7a-hydroxylase activity (3-6).

The molecular mechanism whereby hydrophobic bile salts down-regulate hepatic HMG-CoA reductase activity was not elucidated by previous studies. Some studies have suggested that bile acids down-regulate HMG-CoA reductase directly by demonstrating a reduction in HMG-CoA reductase activity in lymph fistula rats (7). In contrast, other investigators consider the effects of bile acids on cholesterol metabolism to be indirect, i.e. occurring as a result of effect of bile acids on cholesterol absorption (8). HMG-CoA reductase activity ap- pears to be regulated at many levels. It has been demonstrated that HMG-CoA reductase mRNA is regulated at the tran- scriptional level by “oxysterols” which are believed to bind to a repressor protein (9-11). Glucocorticoids have been reported to destabilize hepatic HMG-CoA reductase mRNA in rats (12). In addition, the translation of HMG-CoA reductase mRNA and the turnover of this enzyme has been reported to be regulated both by sterol and by non-sterol derivatives of mevalonate (13-15). Finally, the catalytic efficiency of this enzyme can be altered in vitro via covalent modification (phosphorylation-dephosphorylation) (16) and changes in the lipid composition and fluidity of its membrane microenviron- ment (17). The goal of the current study was to define in greater detail the molecular mechanism(s) by which hydro- phobic bile salts down-regulate HMG-CoA reductase.

EXPERIMENTAL PROCEDURES

Materials Dithiothreitol, bovine serum albumin, leupeptin, EGTA, phenyl-

methylsulfonyl fluoride, proteinase K, ribonuclease A, cholesterol,

9413

9414 Effect of Hydrophobic Bile Ac

NADPH, cholestyramine, and deoxycholic acid were obtained from Sigma. Chenodeoxycholic acid, ursodeoxycholic acid, and cholic acid were purchased from Behring Diagnostics. Mevinolin was a generous gift from Merck Pharmaceuticals. Goat anti-rabbit IgG conjugate and Bio-Rex 5 were obtained from Bio-Rad. The 3,3',5,5'-tetramethyl- benzidine peroxidase system was purchased from Kirkegaard and Perry Laboratories, Inc. (Gaithersburg, MD). The nick translation kit was obtained from Bethesda Research Laboratories and guanidine thiosulfate was purchased from Fluka (Ronkonkoma, NY). Deoxyri- bonuclease I was obtained from Worthington, and the Sephadex G- 60, fine, DNA grade and 3-hydroxy-3-methylglutary-coenzyme A were purchased from Pharmacia LKB Biotechnology Inc. The Im- mulon 1 microwell plates were made by Dynatech (Chantilly, VA). All radioisotopes were purchased from Du Pont-New England Nu- clear. These include ~L-3-[glutaryl-3-'~C]hydroxy-3-methylglutaryl- coenzyme A (57.6 mCi/mmol), RS-[5-3HH-mevalonolactone (38.8 mCi/ mmol)], [a-.'"P]dCTP (3000 Ci/mmol), and [w3'P]GTP (800 Ci/ mmol). All other reagents were of the highest quality available.

Experimental Design Male Sprague-Dawley rats weighing 125-150 g were purchased

from Harlan Sprague-Dawley. The rats were housed in a light- controlled room where the dark period was from 0300 to 1500 h. The animals in experimental groups were fed powdered Purina rodent chow containing 1% (w/w) bile acids (except deoxycholic acid which was fed at 0.5% to reduce the likelihood of hepatotoxicity), or choles- terol (2%) (w/w) for 2 weeks prior to killing. Some rats were fed 5% cholestyramine or 0.1% mevinolin for 5 days prior to killing.

Preparation of Hepatic Microsomes Rats were killed a t 0900 h. The livers were removed and placed in

ice-cold buffer A (100 mM potassium phosphate, pH 7.2, 100 mM sucrose, 50 mM KCI, 1 mM EDTA, 200 mM sodium fluoride, 3 mM dithiothreitol, 100 p~ leupeptin, 5 mM EGTA, 1 mM phenylmethyl- sulfonyl fluoride). Buffer A was changed once to remove blood, the livers were minced, and homogenized by five downward strokes with a motor-driven Teflon pestle in a Potter-Elvehjem glass homogenizer (0.15-mm clearance). The homogenate was centrifuged twice at 10,000 X g for 10 min. The supernatant fluid was collected and then centri- fuged at 100,000 X g for 2 h. All of the above procedures were carried out a t 4 "C. The microsomal pellet was suspended in 10 ml of buffer A and homogenized for five downward strokes. It was then stored in aliquots a t -70 "C. Microsomal protein was quantitated by the method of Lowry et al. (18).

Enzyme Assays

Microsomal HMG-CoA reductase activity was assayed essentially as described by Beg and Stonik (19). The assay was performed in a total volume of 130 p1 a t 37 "C in a 1.5-ml microcentrifuge tube. Microsomes (600 pg) were preincubated for 20 min a t 37 "C with 10 p1 of bovine serum albumin (50 mg/ml) and 10 p1 of a high salt buffer (stock concentration was 1.6 M potassium phosphate, pH 6.9, 2 M KCI, 30 mM EDTA, 2 mM sodium azide, 10 mM dithiothreitol) in a total volume of 100 pl. The reaction was initiated by the addition of 15 pl of a mixture of 2.3 mM NADPH and 174 p M DL-[3-14C]HMG- CoA (4,000-5,000 dpm/nmol). [3H]Mevalonolactone (300,000 dpm/ 15 pl) was added as an internal standard. The reaction was allowed to run for 30 min and then terminated with 50 pl of 10 N HC1. Tubes were centrifuged, and 70-pl aliquots were applied to 1-ml columns of Bio-Rex 5 resin. [:3H]Mevalonolactone was eluted with two 1-ml washes of water. The radioactivity was then determined by liquid scintillation counting.

Preparation of HMG-GOA Reductase Antibodies Microsomes isolated from livers of rats fed 6% cholestyramine for

4 days and killed at the peak of diurnal rhythm were used for solubilization and purification of proteolytically cleaved (53 kDa) electrophoretically homogenous HMG-CoA reductase (20). Antibod- ies to purified rat liver 53-kDa HMG-CoA reductase were prepared in New Zealand White rabbits. Primary immunization of rabbits was achieved by injecting 70 pg of homogenous HMG-CoA reductase in Freund's complete adjuvant. Eight secondary immunizations a t 2- week intervals of 30 pg of HMG-CoA reductase in incomplete adju- vant were followed. Anti-reductase antibody in serum was monitored by immunotritration of enzymic activity. The IgG fractions of both preimmune and immune serum were purified by high performance

:ids on HMG-CoA Reductase

liquid chromatography using a protein A affinity column (21). We have previously established that the antibody preparation used in the current studies is specific both for native membrane hound (97 kDa) and proteolytically cleaved soluble (53 kDa) HMG-CoA reductase fraction (21, 22).

ELISA Assay for Determining HMG-CoA Protein Mass

The assays were performed in 96-well Immulon 1 ELISA plates. Plates were filled with 100 pl of microsomal protein diluted in 0.05 M carbonate-bicarbonate-coating buffer, pH 9.6. This was allowed to sit overnight a t 4 "C. The protein solution was then removed, and the wells were filled with 100 p1 of Blotto (5% non-fat dry milk in Tris- buffered saline). This blocking solution was allowed to incubate for 1 h a t room temperature. The blocking solution was removed and the plates were washed three times, for 2 min each, with 100 p1 of Tween- Tris-buffered saline (TTBS) (20 mM Tris, 500 mM NaC1, pH 7.5, 0.05% Tween-20). The plates were then incubated with 50 pl of rabbit anti-HMG-CoA reductase for 2-4 h a t room temperature. The wells were again washed five times with TTBS. The secondary antibody consisted of 50 pl of goat anti-rabbit IgG conjugated with horseradish peroxidase (1:lOOO). This was allowed to incubate on the plates for 1-2 h. The wells were again washed five times with TTBS. Plates were developed by the addition of 100 pl of the 3,3',5,5'-tetramethyl- benzidine peroxidase system. The color reagent remained on the plates for 30 min, and the development was terminated by the addition of 100 p1 of 1 M phosphoric acid. The absorbance was read at 450 nm on a Bio-Tek model EL 310 automatic microplate reader.

R N A Extraction

The liver was removed and two 1-g pieces taken from different lobes. These pieces were then homogenized separately using Dounce homogenizers in a solution of 4 M guanidine thiocyanate, 10 mM Tris, pH 7.4, and 7% P-mercaptoethanol. Sarcosyl was added to 2%, and the homogenate heated to 65 "C for 5 min and passed through a 23- gauge needle. This suspension was then underlayered with 5.7 M cesium chloride and 10 mM EDTA, pH 7.4. The suspension was centrifuged at 100,000 X g for 16 h. The pellet was washed with ethanol (95%), suspended in distilled water, and precipitated in 95% ethanol a t pH 5.0 and stored a t -20 "C (23).

Isolation of Polyadenylated-RNA

Polyadenylated-RNA (poly(A)+) was isolated from total RNA using an oligo(dT)-cellulose column (1.0-ml volume). Total RNA (1-2 mg/ ml) in 1 mM sodium EDTA, pH 7.0, was heat denatured a t 70 "C for 1 min and cooled on ice. The sample was then added to an equal volume of 2 X binding buffer (1 M lithium chloride, 20 mM Tris-HCI, pH 7.5, 2 mM sodium EDTA and 1% SDS) and applied to the oligo(dT)-cellulose column several times. The column was then washed with 1 X binding buffer until no A26O-absorbing material was detectable in the column eluent. Bound polyadenylated-RNA was eluted with 6 ml of elution buffer (10 mM Tris-HC1, pH 7.5, 1 mM EDTA, and 0.2% SDS). The polyadenylated-RNA was precipitated in 1/10 volume 3 M sodium acetate, pH 5.0, and two volumes of cold 95% ETOH overnight. The polyadenylated-RNA was stored as the ETOH precipitate (-20 "C) until used.

Northern Blot Hybridization Polyadenylated-RNA was size fractionated by electrophoresis in

1% agarose gel containing 7% formaldehyde and then transferred onto nitrocellulose membrane filters overnight capillary blotting as described by Thomas (24). The membranes were baked, prehybridized in 5 X Denhardt's solution, 4 X SSC (standard sodium citrate), 1% SDS, 0.1 mg/ml salmon sperm DNA, and 50% formamide, at 42 "C for 3 h. The solution was changed and the "2P-labeled HMG-CoA reductase cDNA probe (5.5 million cpm/ml) was added. The mem- branes were then hybridized for 16 h at 42 "C. It was washed in 2 X SSC, 0.1% SDS for 5 min a t room temperature, 15 min in 2 X s s c , 0.1% SDS at room temperature, 2 h in 0.1 X SSC, 0.5% SDS a t 50 "C, and 2 times for 30 min each in 0.1 X SSC, 0.5% SDS at 50 "C. The membrane was air dried and analyzed using the same procedures as described for dot blot hybridization. The reprobing of the Northern blot with rat albumin was performed as described for dot blot hybrid- ization (below).

Effect of Hydrophobic Bile Acids on HMG-CoA Reductase 9415

Dot Blot Hybridization

Total RNA was diluted in 20 X SSC, 7.4% formaldehyde and applied to nitrocellulose using a Hybri-Dot Manifold from Bethesda Research Laboratories. The membrane was then baked 1 h at 80 "C in a vacuum oven. The plasmid pRed 227 containing a full-length hamster C-DNA of the HMG-CoA reductase gene (25) was labeled with [a-"'PIdCTP by nick translation. The membrane was prehy- bridized in 4 X SSC, 1% SDS, 5 X Denhardt's, 0.1 mg/ml salmon sperm DNA and 50% formamide for 2 h at 42 "C. The solution was changed and the probe added to 2 million cpm/ml. The membrane was then hybridized for 16 h at 42 'C. It was washed two times in 2 X SSC for 5 min at room temperature, 30 min at 37 "C in 2 X SSC, 0.1% SDS, 30 min at 37 "C in 0.1 X SSC, 0.1% SDS and 30 min at 37 "C in 0.1 X SSC. The membrane was air dried and exposed to x- ray film (XAR) for 24 h at -70 "C. This film was then developed and scanned using a Shimadzu laser densitometer. The membrane was then washed three times in boiling water for 5 min in order to remove the HMG-CoA reductase probe.

The insert of plasmid pRSA 57 (26), containing a rat albumin C- DNA, was then labeled with [a3*P]dCTP as above, and the same prehybridization and hybridization conditions were used. The mem- brane was then washed at room temperature using the same solutions and times. It was air dried and exposed for 2 h at -70 "C, and the dots cut out and counted by liquid scintillation spectrometry.

The optical densities for pRed-227 and the cpm/dot for pRSA 57 were plotted separately and the slopes calculated. The pRed-227 slope was divided by the slope for pRSA 57 to obtain an index for each condition and the resulting indices compared to the control to deter- mine the percent of control.

Nuclear Run-on Studies Isolation of Nuclei-One g of rat liver was minced in 0.32 M sucrose,

:3 mM MgC12, 1 mM HEPES, pH 6.8, a t 4 "C and homogenized in a motor-driven Teflon/glass homogenizer (0.15 mm clearance) a t 500 rpm for two strokes at 4 "C. The homogenate was then centrifuged at 700 X g for 10 min at 4 "C. The pellet was resuspended in cold 2.1 M sucrose, 1 mM MgC12, 1 mM HEPES, pH 6.8, and centrifuged at 50,000 X g for 80 min at 4 "C. The nuclei were then washed in 0.25 M sucrose, 1 mM MgC12, 1 mM HEPES, pH 6.8, and suspended in 20 mM Tris-HC1, pH 7.9, 30% glycerol, 140 mM KCl, 5 mM MgC12, 1 mM MnCl?, and 14 mM p-mercaptoethanol. The nuclei were counted and aliquoted at 4 X 107/200 p1 and frozen at -70 "C until used (27).

RNA Labeling and Isolation-An aliquot of frozen nuclei was added to 200 p1 of 10 mM Tris-HC1, pH 8.0, 5 mM MgCl,, 0.3 M KCl, 1.25 mM ATP, 1.25 mM CTP, 1.25 mM UTP, 5 mM dithiothreitol, and 30 pl of [a-:"P]GTP (800 Ci/mmol, 10 mCi/ml) was added to each. They were incubated for 30 min at 30 'C, and the reaction stopped by the addition of 40 pg of DNase I in 0.5 M NaCl, 50 mM MgCL, 2 mM CaC12, 10 mM Tris-HC1, pH 7.4. After 5 min at 30 'C, 200 pg of proteinase K in 0.5 M Tris-HC1, pH 7.4,5% SDS, 0.125 M EDTA was added and incubated 45 min at 42 "C. This solution was then extracted with 1 ml of phenol/chloroform/isoamyl alcohol (25:241 v/v/v) and the interface was re-extracted with 1 ml of 10 mM Tris-HC1, pH 7.4, 1 mM EDTA (TE buffer). The combined aqueous phases were precip- itated with 1/10 volume of 3 M sodium acetate and two volumes of ethanol using a dry ice/ethanol bath for 30 min. The RNA was recovered by centrifugation at 10,000 X g for 30 min and resuspended in 100 pl of TE buffer. This was then applied to a spin-column made from a 1-ml syringe and Sephadex G-50, fine, DNA grade. The column was centrifuged 2 min at 200 X g and washed twice with 100 pl of TE buffer. The RNA was mildly digested by adding Vi volume of 1 M NaOH on ice for 5 min and neutralized by adding Yz volume of 1 M HEPES. This was precipitated with 2.5 volumes of ethanol at -20 "C overnight (28).

Hybridization-Nitrocellulose strips containing spots of dilutions of pRed-227 and pRSA 57 were placed in small heat-sealable bags with 4 ml of 10 mM TES, pH 7.4, 10 mM EDTA, 0.2% SDS, 0.3 M NaCI, and 1 X Denhardt's (hybridization solution). They were pre- hybridized for at least 2 h at 65 "C. The RNA was recovered by centrifugation at 4 "C and resuspended in 100 pl of 10 mM TES, pH 7.4, 10 mM EDTA, 0.2% SDS. An aliquot was counted, the samples were standardized to the lowest counts, and the appropriate volume added to fresh hybridization buffer in each bag. Hybridization was performed at 65 "C for 44 h. The strips were rinsed 5 min at 65 "C in 2 X SSC and washed two times for 1 h at 65 "C in 2 X SSC. They were then incubated at 37 "C for 30 min in 40 ml of 2 X SSC containing 400 pg of RNase A, washed 1 h at 37 "C in 2 X SSC and

then air dried. They were exposed to x-ray film (XAR) for 3-10 days (29).

Statistical Analysis-Data are reported as mean * S.D. A two- tailed T test for unpaired samples was used to determine statistical differences. These were determined by using a SAS statistical package (SAS Institute, Inc., Cary, NC) on an IBM PS2 model 80 computer.

RESULTS

Measurement of HMG-CoA Reductase Activity and Protein Mass-In the current study, rats were fed either bile acids of varying hydrophobicity or cholesterol which was added to laboratory chow. Livers from these animals were harvested, divided in half, and total RNA extracted or microsomes prepared as described under "Experimental Procedures." The data in Fig. 1 show the specific activities (solid bars) of HMG- CoA reductase in rat liver microsomes prepared in groups of animals. Ursodeoxycholate, a representative hydrophilic bile acid, decreased HMG-CoA reductase activity only minimally. In contrast, more hydrophobic bile acids, e.g. cholate, cheno- deoxycholate (fed as 1% w/w), and deoxycholate (fed as 0.5% w/w) decreased (57-77%) enzyme specific activity; this de- crease was statistically significant ( p < 0.025). As expected cholesterol feeding decreased specific activity by 89% of con- trol value. Deoxycholic acid was fed at 0.5% to avoid toxicity.

We next sought to quantitate the effects of bile acid and cholesterol feeding on HMG-CoA reductase protein mass using a newly developed ELISA method described under "Experimental Procedures." Using rabbit polyclonal antibody raised to purified HMG-CoA reductase, we were able to detect as little as 0.8 ng of pure protein (53-kDa fragment). The data in Fig. 2 show a representative ELISA assay of HMG-CoA reductase protein under varying experimental conditions. Se- rial dilutions of microsomal protein from 1 to 0.125 pg were used. As compared to control, cholestyramine feeding caused the expected increase in HMG-CoA reductase protein, while cholesterol feeding caused a decrease. UDCA caused only a slight decrease in HMG-CoA reductase protein as compared to control. In contrast, chenodeoxycholate and deoxycholic acid caused a significant decrease in HMG-CoA reductase protein similar to that seen with cholesterol feeding.

Both HMG-CoA reductase specific activity (solid bars) and protein mass (stippled bars) from various groups of animals are shown together in Fig. 1. The hydrophobic bile acids

1w y.03

&a

Per Cent of w

Control

40

20

n 1% u r n

N=5 l X C A IXCDCA N=5 N=5

0.5% DCA 2% Cholesterol N=5 N=5

FIG. 1. Effect of bile acid or cholesterol feeding on hepatic HMG-CoA reductase specific activity and protein mass. Ur- sodeoxycholic acid (UDCA), cholic acid (CA), chenodeoxycholic acid (CDCA), deoxycholic acid (DCA), or cholesterol (rol) were fed to animals for 14 days, hepatic microsomes prepared, and specific ac- tivities (solid bars) and protein mass (cross-hatch burs) determined as described under "Experimental Procedures." The control specific activity (100%) was 17.8 pmol/min/mg protein. Protein mass was quantitated over a wide (0.125-1 pg) of microsomal protein (Fig. 2). N. = number of experimental animals and arrow bars are * S.D.

9416 Effect of Hydrophobic Bile Acids on HMG-CoA Reductase

A 5% Cholestyramine A 1% CDCA 0 2% Cholesterol

0 In -2- 0.”

u C 0

0 cn D

0 . a

0.2w

0 2w 400 6w ” 1 w O

Microsomal Protein (ng)

PI(;. 2. Quantitation of microsomal HMG-CoA reductase p r o t e i n mass under different experimental conditions. Micro- s(1111~s \yere prepared from livers n ! animals fed either hile acids, cholesterol or cholestyramine. and reductase protein quantitated over a \vide ranxe (0.125-1 p g ) o f microsomal protein as described under “Esl,erimental I’roceclures.”

significantly reduced HMG-CoA reductase protein mass; cho- late by (27r;), chenodeoxycholate by (31761, and deoxycholate ( 4 2 5 ) . These differences were statistically significant for all three hydrophobic bile acids ( p < 0.025 t o 0.0001) when compared to controls. Ursodeoxycholate feeding reduced pro- tein mass by only 16%; this decrease was not. statistically significant. The amount of HMG-CoA reductase protein was reduced by 3 5 % in cholesterol-fed animals. In all groups tested, the fractional changes observed in protein mass are significantly !ess than we observed in specific enzyme activity (I-, < 0.0001 by paired t test).

Mmsurement of HMG-CoA Reductase mRNA Levels-We next quantitated the levels of HMG-CoA reductase mRNA in the various groups of animals. Northern blot analysis was carried out on poly(A)+ mRNA isolated from animals fed mevinolin (induced), cholestyramine (induced), various bile acids and cholesterol (repressed) (Fig. 3A). The membrane filters were reprobed with albumin c-DNA which served as a n internal control (Fig. 3 R ) . Northern blot analysis of total RNA from these samples showed similar relative concentra- tions as poly(A)+ mRNA (data not shown). In order to more accurately quantitate HMG-CoA reductase mRNA, serial di- lutions of total RNA were dot-blotted onto nitrocellulose memhrane filters and hybridized first with the pRED-227 plasmid as described under “Experimental Procedures.” This plasmid contains a full-length hamster c-DNA from the HMG-CoA reductase gene. The amount of hybridization was quantitated by laser densitometry. The HMG-CoA reductase probe was then st.ripped from the blots and the filters rehy- bridized with a rat serum albumin probe which served as a n internal control.

There was a marked decrease in levels of HMG-CoA reduc- tase mRNA in livers prepared from rats fed relatively hydro- phobic bile acids (Fig. 4, solid bars). Cholate, chenodeoxycho- late, and deoxycholate reduced HMG-CoA reductase mRNA by 43, 47, and 54%, respectively. Cholesterol feeding reduced HMG-CoA reductase by 53%. In contrast, ursodeoxycholate reduced HMG-CoA reductase mRNA levels by only 10%.

Nuclear Run-on Assays for HMG-CoA Reductase Transcrip- tional ActiuitF-Livers were harvested in animals fed the various diets, divided in half, and nuclei and microsomes prepared as described under “Experimental Procedures.’’ The amounts of labeled reductase and albumin mRNA were quan- titated by laser densitometry and the ratio of reductase to alhumin mRNA was determined. These ratios from different

B

?X s

I R s

FIG. 3. Hepatic HMG-CoA reductase mRNA levels under various feeding conditions. This Northern hlot (panrl A ) com- pares reductase poly(A)’ mRNA levels in animals led mevinolin (lonc I ) , cholestyramine (lanc 2). control chow (lane 3 ) . ursodeoxycholic acid (lane. 4 ) . cholic acid (lane 5 ) , chenodeoxycholic acid ( l a w 6) . deoxycholic acid (lane 7), and cholesterol (lanc 8) as descrihed under “Experimental Procedures.” Polg(A)+ RNA (5 pg/lane) was isolated from liver and separated hv size on a ICE agarose gel containing formaldehyde. RNAs were then transferred to nitrocellulose mem- hranes and hvbridized with ‘”P-laheled pRED 227 cDNA. The mem- brane shown in pond A was “stripped” of lahel and hyhridized with .‘T-laheled pRSA 57 alhumin c-DNA (panrl H ) .

175

150 I 1

IXUDCA I%CA 1PICDCA 0 SXDCA Chome?lerol

N=4.4 N=4.4 N=4.4 N=4.4 N 5 4 . 2

FIG. 4. Effect of bile acid or cholesterol feeding on steady state mRNA levels and relative transcriptional rates of he- patic HMG-CoA reductase. The relative amounts of steady state mRNA (dark bars) are shown for the various feeding conditions. Reductase mRNA and transcriptional rates (cross-hatch bars) are normalized to that of alhumin as descrihed under “Experimental Procedures.” N = numher of experimental animals and error bars are +. S.E. See Fig. 1 for abhreviat.ions of hile acids. N S , -.

feedings were compared to control and the relative values are shown in Fig. 4 (stippled bars). Transcriptional activity in nuclei from CA and cholesterol fed animals was much lower (46-51%) than control. Surprisingly, the transcriptional ac- tivities of CDCA and DCA-fed animals were consistently near or above control values (Fig. 4, stippled bars), even though the

Effect of Hydrophobic Bile Acids on HMG-CoA Reductase 9417

steady state mRNA levels were markedly reduced (Fig. 4, dark bars).

Time Course of HMG-CoA Reductase Activity, Protein Mass, and mRNA during Feeding of Chenodeoxycholate-Cheno- deoxycholic acid (1% w/w) was fed to rats over a 14-day time course. Livers were prepared from animals on day 2, 4, 8, 11, and 14, and the relative levels of HMG-CoA reductase activ- ity, protein, and mRNA were determined as described under “Experimental Procedures.” HMG-CoA reductase-specific ac- tivity dropped 43% in 2 days whereas protein mass decreased 21%. The relative amount of HMG-CoA reductase mRNA decreased 58% in 2 days. There was a slow decline in all three measured parameters over the next 12 days (Fig. 5 ) .

DISCUSSION

Hepatic HMG-CoA reductase specific activity can poten- tially be regulated by many different mechanisms. Reductase catalytic activity can be altered in vitro by a phosphorylation- dephosphorylation control system. Phosphorylation, which inhibits catalytic activity, can be carried out by a t least three different protein kinases (16). In addition, Davis and Poznan- sky (17) showed that reductase activity can be altered by changes in membrane fluidity. An increase in membrane fluidity in human skin fibroblast was associated with an increase in reductase activity. Transcriptional control of re- ductase gene activity has been hypothesized to be under end- product repression control by oxysterols (9-11). The oxyste- rols which have been reported to increase in cholesterol fed animals include 24-, 25- , and 26-hydroxycholesterol (30). These compounds are postulated to bind to a cytosolic recep- tor which translocates to the nucleus and “down-regulates” reductase gene transcriptional activity. Alternatively, these sterols may induce a nuclear protein that binds to the sterol regulatory element of the promoter of HMG-CoA reductase (31). The degradation of reductase protein can be accelerated by sterols and/or non-sterol mevalonate metabolites ( 2 5 , 32). Finally, Simonet and Ness (12) have reported that glucocor- ticoids can markedly decrease the half-life of HMG-CoA reductase mRNA in hypophysectomized rats. Moreover, Au- werx et al. (33) demonstrated changes in HMS-CoA reductase mRNA stability in human monocytic leukemia cell lines by stimulators of protein kinase C.

In our previous studies, we showed that both reductase and cholesterol 7a-hydroxylase activity decreased following feed- ing of relatively hydrophobic, but not hydrophilic, bile acids (3-6). We observed a highly significantly inverse linear rela- tionship between the bile acid pool hydrophobicity index and the specific activities of these two enzymes. Hydrophobic bile acids affected reductase activity in the order of increasing

I 0 2 4 6 8 1 0 1 2 1 4

Time (days)

FIG. 5. Time course of reductase specific activity, protein mass, and mRNA in animals fed chenodeoxycholic acid. Quan- titation of‘ reductase specific activity. protein mass, and mRNA are described under “Experimental Procedures.”

hydrophobicity (CA < CDCA < DCA). In the present study, we show a highly significant and reproducible decrease in reductase specific activity (Fig. 1) and steady state mRNA levels (Fig. 4) in livers from animals fed relatively hydrophobic bile acids, cholic acid, chenodeoxycholic acid and deoxycholic acid. In contrast, ursodeoxycholic acid, a relatively hydro- philic bile acid, affected reductase activity, and mRNA to only a small extent.

The fractional decrease in reductase protein mass (Fig. 1) following feeding of relatively hydrophobic bile acids was significantly less prominent than the changes in specific ac- tivity (Fig. 1) or steady state mRNA levels (Fig. 4). This could be due to cross-reactivity of the antibody preparation to other microsomal antigens or due to presence of a substantial amount of inactive reductase protein. Consistent with earlier results (22), Western blot analysis showed that antibodies to purified reductase (53,000-dalton fragment) recognized only a single detectable protein (94,000 daltons) in microsomes prepared from cholestyramine-fed animals. In addition, the preparation of rat liver microsomes in the presence and ab- sence of sodium fluoride did not change the amount of im- munoreactive protein in ELISA assays, suggesting that under our assay conditions, the state of phosphorylation of the enzyme does not affect immunoreactivity (20-22). We believe that there may be a considerable amount of inactive reductase protein in microsomes. Several laboratories have reported the presence of inactive forms of HMG-CoA reductase which may be present in vivo (34-36). We previously purified and char- acterized an inactive form of HMG-CoA reductase from rat liver that differed from the active enzyme by a single peptide (37). There was approximately three times the amount of inactive HMG-CoA reductase isolated by HMG-CoA affinity chromatography, as compared to active enzyme (37). These results are consistent with the high “background observed in our ELISA assays (Fig. I). It is not certain whether this inactive protein represents a pool of enzyme precursor (a proenzyme), covalently modified reductase, or an artifact generated during the prepdration of microsomes.

We have shown that feeding cholic acid, but not cheno- deoxycholic acid or deoxycholic acid, markedly reduces reduc- tase transcriptional activity (Fig. 4). We hypothesize that cholic acid down-regulates reductase activity both directly and indirectly. A number of studies in the rat (38-42) report that cholic acid is a much more powerful promotor of cholesterol absorption than chenodeoxycholic acid or deoxycholic acid, although the latter are more potent detergents. This would result in enhanced delivery of cholesterol to the liver via chylomicron remnant cholesterol causing enhanced end-prod- uct repression of reductase gene transcriptional activity. Pre- liminary data in the chronic bile fistula rat model shows that intravenous infusion of cholic acid decreases HMG-CoA re- ductase activity.’ In this model, intestinal cholesterol absorp- tion does not occur suggesting that cholic acid may also play a direct role in the regulation of hepatic HMG-CoA reductase.

The most surprising result of the current study was the different transcriptional rates observed in hepatic nuclei iso- lated from cholic acid as compared to chenodeoxycholic acid and deoxycholic acid-fed rats (Fig. 4). In contrast to cholic acid, the relative transcriptional rates in liver nuclei isolated from chenodeoxycholic acid or deoxycholic acid-fed animals were near control levels, but steady state mRNA levels were near fully repressed levels. We hypothesize that these hydro- phobic bile acids post-transcriptionally decrease HMG-CoA reductase mRNA levels. In this regard, Simonet and Ness (12) have previously demonstrated a marked decrease in

. .-

’ P. F. Duckworth, unpublished results.

9418 Effect of Hydrophobic Bile Acids on HMG-CoA Reductase

HMG-CoA reductase mRNA half-life in hypophysectomized 20. Beg, Z. H., Stonik, J. A., and Brewer, H. B., Jr. (1980) J. Biol. rats treated with glucocorticoids. We believe that hydrophobic Chem. 266,8541-8545 bile acids may be important physiological regulators of cho- 21. Beg, Z. H., Stonik, J . A., and Brewer, H. B., Jr. (1987) J. Biol.

and degradation must be tightly regulated. Chem. 260,1682-1687 lesterol synthesis in the liver, where the pathways of synthesis 22. Beg, Z. H., Stonik, J. A., and Brewer, H. B., Jr. (1985) J , Bioi,

Chem. 262,13228-13240

REFERENCES

1. Danielsson, H., Einarsson, K., and Johansson, G. (1967) Eur. J.

2. Bjorkhem, I. (1985) in Sterols and Bile Acids (Danielsson, H., and

23. Chirgwin, J. M., Przybyla, A. E., MacDonald, R. J., and Rutler,

24. Thomas, P. S. (1980) Proc. Natl. Acad. Sci. U. S. A. 77, 5201-

25. Chin, D. J., Gil, G., Faust, J. R., Goldstein, J. L., Brown, M. S.,

N. J. (1979) biochemistry 18,5294-5299

5205

and Luskey, K. L. (1985) Mol. Cell Biol. 5,634-641

Y ork A. A., and Bonner, J. (1979) Proc. Natl. A c d . Sci. U. S. A. 76,

M.* Hartman, '., and V1ahcevic~ z. R. (lgg8) Hepatolog3' '9 27. Clarke, C. F., Fogelman, A. M., and Edwards, P. A. (1985) J. Bwl.

Biochem. 2,44-49

Sjovall? Jv eds) PP. 231-278, Science New 26. Sargent, T. D., Wu, J., Sala-Trepat, J. M., Wallace, R. B., Reyes,

3. Heuman, D. M., Hernandez, C. R., Hylemon, P. B., Kubaska, W. 3256-3260

358-365 4. Heuman, D. M.9 Vlahcevic, z. R.9 Bailey, M. L.7 and HYlemon, p. 28. Greenberg, M., and Bender, T. (1990) in Current Protocok in

Chem. 260,14363-14367

Molecular Biology (Ausubel, F., et al., eds) pp. 4.10.1-4.10.9, Wiley-Interscience, New York

B. (1988) Hepatology 8, 892-897 5. Heuman, D. M. (1988) J. Lipid Res. 30, 719-730 6. Heuman, D. M v HYlemon, p. B.7 and Vlahcevic, z. R. (1989) J. 29. Linial, M., Gunderson, N., and Groudine, M. (19%) Science 230,

Lipid Res. 30, 1161-1171 7. HamPrecht, B.7 Roscher, R.9 Waltinger, G., and Nussler, c. (1971) 30. Saucier, S. E., Kandutsch, A. A., Gayen, A. K., Swahn, D. K.,

Eur. J. Biochem. 18, 15-19 and Spencer, T. A. (1989) J. Bwl. Chem. 264,6863-6869 8. SPadY, D. K., Stan& E. F., Bilhafiz, L. E., and DietschY, J. M. 31. Rajavashisth, T. B., Taylor, A. K., Andalibi, A., Sevenson, K. L.,

(1986) Proc. Natl. Acad. Sci. U. S. A. 83, 1916-1920 and Lusis, A. J. (1989) Science 246,640-643 9. Osborne, T. F.7 Gil, G.9 Goldstein, J. L., and Brown, M. s. (1988) 32. Edwards, P. A., Lan, S. F., Tanaka, R. D., and Fogelman, A. M.

J. Biol. Chem. 263,3380-3387 (1983) J. Biol. Chem. 258,7272-7275 10. Dawson, p. A., Van Der Westhuyzen, D. R., Goldstein, J. L., and 33. Auwerx, J. H., Chait, A., and Deeb, S. S. (1989) Proc. Natl. Acad.

Brown, M. S. (1989) J. Biol. Chem. 264, 9046-9052 Sci. U. S. A. 86, 1133-1137 11. Saucier, s. E., Kandutsch, A. A., Phima, S., and Spencer, T. A. 34. Erickson, S. K., Shrawsburg, M. A., and Gould, R. G. (1980)

(1987) J. Biol. Chem. 262,14056-14062 Biochim. Biophys. Acta 620, 70-79

573 Natl. Acad. Sci. U. S. A. 77,6429-6433

1126-1132

12. Simonet, W. s., and Ness, G. c. (1989) J. Biol. Chem. 264, 569- 35. Arebalo, R. E., Hardgrave, J. E., and Roland, B. J. (1980) Proc.

13. Brown, M. S., and Goldstein, J. L. (1980) J. Lipid Res. 21,515- 36. Arebalo, R. E., Tormanen, C. C., and Hardgrave, J. E. (1982)

14. Nakanishi, M., Goldstein, J. L., and Brown, M. S. (1988) J. Biol. 37. Whitehead, T. R., Vlahcevic, Z. R., Beg, Z. H., and Hylemon, P.

15. Peffley, D., and Sinensky, M. (1985) J. Biol. Chem. 260, 9949- 38. Watt, S. M., and Simmonds, W. J. (1984) J. Lipid Res. 26, 448-

16. Beg, Z. H., Stonik, J. A., and Brewer, H. B., Jr. (1987) Metabolism 39. Raicht, R., Cohen, B. I., and Mosbach, E. H. (1974) Gmtroenter-

17. Davis, P. J., and Poznansky, M. J. (1987) Proc. Natl. Acad. Sci. 40. Cohen, B. I., Raicht, R. F., and Mosbach, E. H. (1977) J. Lipid

18. Lowry, 0. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. 41. Swell, L., Flick, D. F., Field, H., and Treadwell, C. R. (1953) Proc.

19. Beg, Z. H., and Stonik, J. A. (1982) Biochem. Biophys. Res. 42. Vahouny, G. V., Gregorian, H. M., and Treadwell, C. R. (1959)

517 Proc. Natl. A c d . Sci. U. S. A. 79, 51-55

Chem. 263,8929-8937 B. (1984) Arch. Biochem. Biophys. 230,483-491

9952 455

36,900-917 Ology 67, 1155-1161

U. S. A. 84,118-121 Res. 18,223-231

(1951) J. Eiol. Chem. 193,265-275 SOC. Exp. Biol. Med. 84, 428-431

Commun. 108,559-566 Proc. SOC. Enp. Biol. Med. 101, 538-540