the active site of hamster 3-hydroxy-3-methylglutaryl-coa

THE JOURNAL OF BIOWGICAL CHEMISTRY 0 1994 by The American Society for Biochemistry and Molecular Biology, Inc

Vol. 269, No. Issue of January 14, PP . 1217-1221, 1994 Printed in U.S.A.

The Active Site of Hamster 3-Hydroxy-3-methylglutaryl-CoA Reductase Resides at the Subunit Interface and Incorporates Catalytically Essential Acidic Residues from Separate Polypeptides*

(Received for publication, June 23, 1993)

Kenneth Frimpong and Victor W. Rodwelll From the Department of Biochemistry, Purdue University, West Lafayette, Zndiana 47907-1153

We employed site-directed mutagenesis based on sequence comparisons and characterization of purified mutant enzymes to identify Gld6* and Asp7= of Syrian hamster 3-hydroxy-3-methylglutaryl coenzyme A (HMG- CoA) reductase (EC 1.1.1.34) as essential for catalysis. Mutant enzymes E558D, E558Q, and D766N had wild- type K, values for (S)-HMG-CoA and NADPH, but exhibited less than 0.5% of the wild-type catalytic activity. The inactive mutant polypeptides E558Q and D766N never- theless can associate to generate an active enzyme. In vitro, 6% of the wild-type activity was observed when mutant polypeptides E558D and D766N were mixed in the absence of chaotropic agents. When mutant polypeptides E558Q and D766N were co-expressed in Esch- erichia coli, the resulting purified enzyme had 25% of wild-type activity. Hamster HMG-CoA reductase thus is a two-site, dimeric enzyme whose subunits associate to form an active site in which each monomer contributes at least one residue (e.g. G1u568 from one monomer and Asp7= from the other). The wild-type enzyme behaves as a dimer during size exclusion chromatography and has one HMG-CoA binding site per monomer. Syrian hamster HMG-CoA reductase thus appears to be a homodimer with two active sites which are located at the subunit interface.

3-Hydroxy-3-methylglutaryl coenzyme A (HMG-CoAIl reductase (EC 1.1.1.34), which catalyzes the reaction,

HMG-CoA + 2 NADPH + 2 H’ -, mevalonate + 2 NADP+ + CoASH

is rate-limiting for mammalian cholesterol biosynthesis, and therefore constitutes the target enzyme of many drugs aimed at lowering the rate of cholesterol biosynthesis (1).

Guided by the mechanism proposed by Veloso et al. (21, we previously identified Glus3 (3) and His3s1 (4) of the Pseudomo- nas mevalonii enzyme as important for catalysis. Mechanistic investigations of HMG-CoA reductases of higher eukaryotes have long been hampered by the small quantities present in tissues and by the heterogeneity of the purified enzyme, whether derived from tissues or from the overexpressed catalytic domain of the human (5) or radish (6) enzyme. These difficulties have been overcome by the construction of an expression vector derived from pT7-7 which allows purification of

* This work was funded by National Institutes of Health Grant HL 47113. The data derive from the Ph.D. thesis material of K. Frimpong. This is paper 13829 from the Purdue University Agricultural Experi- ment Station. The costs of publication of this article were defrayed in

hereby marked “aduertisement” in accordance with 18 U.S.C. Section part by the payment of page charges. This article must therefore be

1734 solely to indicate this fact. $ To whom requests for reprints should be addressed.

The following abbreviations are used: HMG, 3-hydroxy-3-methylglutaryl; PAGE, polyacrylamide gel electrophoresis.

the catalytic domain of hamster HMG-CoA reductase to a homogeneous state in high yield (7). This development led to the identification of Hiss65, which corresponds to His381 of the €? mevalonii enzyme, as the only histidine of the catalytic domain of the hamster enzyme important for catalysis (8).

While most investigators agree that HMG-CoA reductase is functional as a multimeric entity, no clear consensus has emerged concerning the number of subunits which constitute a functional catalytic unit. Based on gel filtration data, the catalytic domain of the human enzyme appears to be a dimer ( 3 , while the catalytic domains of rat liver (9, 10) and yeast (11) HMG-CoA reductases are said to be tetrameric. Based on radiation inactivation data, the membrane-bound holoenzyme form of the rat liver enzyme is, however, reported to be dimeric (12). The only bacterial HMG-CoA reductase, that from P. mevalonii, was initially reported to be a tetramer (131, although crystallographic studies reveal the presence in the crystal of a trimer of dimers2 with inter-subunit binding sites for substrates (14).

We here identify 2 acidic residues important for catalysis by Syrian hamster HMG-CoA reductase, and show that the active enzyme is an a2 homodimer with its catalytic site at the subunit interface.

MATERIALS AND METHODS Chemicals-Purchased reagents included blue Sepharose (Pharma-

cia LKL? Biotechnology Inc.); T4 DNA ligase and a SequenaseR kit (U. S. Biochemical Corp.); restriction enzymes (New England Biolabs, Pro- mega, or U. S. Biochemical Corp.); p-mercaptoethanol (Sigma), [cY-~~SI~ATP (Amersham Corp.); BioLyte 6-8 ampholyte, BioLyte 3-10 ampholyte, and molecular weight standards for gel filtration and for SDS-PAGE (Bio-Rad); urea (Life Technologies Inc.); Nonidet P-40 (Gal- lard Schlesinger); and Centricon-30 filtration apparatus (Amicon). En- zymically synthesized (S)-HMG-CoA (15) was 97% homogeneous as judged by high performance liquid chromatography (16). All other materials were from previously cited sources (3, 4, 17). Buffer A contained 50 m~ NaCl, 10% (v/v) glycerol, 100 mM sucrose, 10 mM dithiothreitol, and 20 m~ Na,PO,, pH 6.7 or 7.3.

DNA Sequencing-The Sanger dideoxynucleotide chain termination method (18) used for sequencing employed a US. Biochemicals Se- quenaseR kit and [d5S1dATP. Double-stranded sequencing was conducted according to Hsiao (19). Subsequent procedures followed the instructions provided with the SequenaseR kit.

Vectors andBacteria1 Strains-Vector M13mp18-RC,, (8), pACYC-177 (20), pT7-7 (21), pKFT7-2, and pKFT7-21 (7) have been previously described. Escherichia coli TG1 cells (22) served as hosts during mutagenesis and for plasmid preparations. E. coli strain SP1515, a r e d - derivative of strain BL21(DE3) (21), was a generous gift from Ihor Skrypka and Ronald Somerville.

Media for Cell Growth-Cells transformed with either pT7-7 or pKFT7-21 vectors were grown in LB medium (23) containing 50 pg/ml ampicillin. Cells transformed with pACYC-177-based vectors were grown in LB medium (23) containing 20 pg/ml kanamycin. SP1515 cells

C. M. Lawrence and C. V. Stauffacher, unpublished observations.

1217

1218 Active Site of Hamster HMG-CoA Reductase

Wild-type 3. D652N

TABLE I Mutant oligonucleotides

E558Q, D652N, and D766N. The altered residue is shown in bold. Shown are the oligonucleotides that encode mutant enzymes E558D,

Altered bases are underlined. Oligonucleotides 1 and 2 correspond to the coding strand, while oligonucleotides 3 and 4 correspond to the noncoding strand.

Enzyme Oligonucleotide

560 559 558 557 556 Cys Gly Glu T h r T h r

Wild-type 1. E558D 2. E5586 5"AG ACA GCC TTG CGT TGT TG-3'

5"AG ACA GCC TTC CGT TGT TG-3' 5"AG ACA GCC GTC CGT TGT TG-3'

-

649 650 651 652 653 654 L y s T h r Gly Asp Ala Met

5"AAG ACG GGG GAT GCC ATG G-3' 5"AAG ACG GGG F T GCC ATG G-3'

763 764 165 7 6 6 1 6 1 7 6 8 Cys Gly Gln Asp Ala Ala

Wild-type 4. D766N

5"TGT GGC CAG GAT GCA GCA C-3' 5"TGT GGC CAG S T GCA GCA C-3'

transformed with both pT7-7-based and pACYC-177-based vectors were grown in LB medium containing 50 pg/ml ampicillin and 20 pg/ml kanamycin.

Assay of HMG-CoA Reductase ActiuityThe HMG-CoA-dependent decrease in optical density a t 340 nm which accompanied the oxidation of NADPH was monitored in a Hewlett-Packard model 8452 diode array spectrophotometer equipped with a cell holder maintained at 37 "C. Assays were conducted in 150 pl of 100 mM NaCI, 10 mM dithiothreitol, 1.0 mM EDTA, in 100 mM Na,PO.,, pH 6.75. Standard reactant concentrations were 32 p (S)-HMG-CoA and 270 PM NADPH. One enzyme unit corresponds to the oxidation of 1 nmol of NADPH (0.5 nmol of (S)-HMG-CoA) per min. Protein concentration was determined by the method of Bradford (24) using bovine serum albumin as standard.

Site-directed Mutagenesis-Mutagenic oligonucleotides synthesized at the Purdue University Laboratory for Macromolecular Structure were purified by thin-layer chromatography using a U. S. Biochemicals SurePureR oligonucleotide purification kit. Mutations employed an Am- ersham oligonucleotide-directed mutagenesis system (22) and phage or phagemid template prepared as described by Promega (25). All mutations were verified by double-stranded sequencing. Oligonucleotides 1 and 2 (Table I) were annealed to M13mp18-Rc,, to create the indicated mutations at position 558. The mutated genes were inserted into pKFT7-2, then into p m 7 - 2 1 for verification of mutations and for protein expression. Oligonucleotides 3 and 4 were used to create, di- rectly in pKFT7-21, the indicated mutations at positions 652 and 766 (Table I).

Expression and Purification of Mutant Enzymes-Wild-type and mutant enzymes were expressed behind the T7 promoter of pKFT7-21, pACT7, or pT7-7. E. coli strain SP1515 harboring expression vectors was grown a t 37 "C, with shaking a t 300 rpm, in LB medium containing 50 pg/ml ampicillin, 20 pg/ml kanamycin, or both antibiotics where appropriate. Purification of wild-type and mutant enzymes was conducted as previously described for the wild-type enzyme (7).

RESULTS Sequence Alignments-Computer-assisted alignment of the

DNA-derived primary structures of the catalytic domains of 15 HMG-CoA reductases revealed 4 conserved acidic residues. For the hamster enzyme, these are Glu55s and Asp766 (Fig. l) ,

and Asp652. G ~ u ~ ~ ~ corresponds to G ~ u ~ ~ of the l? mevalonii enzyme (3). Since l? mevalonii mutant enzyme E52Q exhibited a severely impaired capacity to bind substrates (31, G ~ u ~ ~ ~ of the hamster enzyme was not further investigated.

Overexpression of Wild-type and Mutant Enzymes-We employed site-directed mutagenesis to alter codon 558 to Asp and Gln, and codons 652 and 766 to Asn. Each mutant gene was then overexpressed in E. coli. Mutant enzyme D652N was both partially degraded and insoluble, and was not further investigated. The soluble mutant enzymes E558D, E558Q, and D766N were purified to over 95% homogeneity as judged by SDS-PAGE

Hamster (27,28) Human (29) Rat Xenopus laevis (30) S e a urchin (31) Drosophila melanogaster (32) Schistosoma mamoni (33) Yeast I & 2 (34.35) Potato 47 Arabidopsis thaliana (36,37) Potato 17 Tomato 2 &. 3 Hevea brasiliensis (38) Haloferax volcanii (39). Pseudomom mevalonrr (40) Consensus

V P M A T T V P M A T T

V P M A T T V P M A T T V P M A T T

~ P M A T T

E P ~ A T T pi%:: V P M A T T V P M A T T V P M A T T F P ~ A T T v P L m V P L A T T

8 G C L V A G C L V A G C L V A G C L V A G C L V A

G C L V A G C L V A G C L V A G C L V A G C L V A G C L V A G C L V.A

m V A G A L L A

G C L V A

C ~ L V A

FIG. 1. Amino acids of the catalytic domains of HMGCoA reductases adjacent to Glum and Asp788 of the Syrian hamster enzyme. Alignments were produced by the Pileup program of the Wis- consin package (26). Sequences generously provided prior to their publication by the listed investigators included tomato (Jonathon Narita and Willie Gruissem, Dept. of Plant Pathology, University of California, Berkeley), potato (Doi Choi and Richard M. Bostock, Dept. of Plant Pathology, University of California, Davis, CA), and rat (David J. Sha- piro, University of Illinois, Urbana, IL).

96 66 - 45 -.- w u v r w

36

21 FIG. 2. SDSpolyacrylamide gel electrophoresis of purified

wild-type and mutant enzymes. Samples in 1% SDS, 50 mM dithiothreitol, 10% (v/v) glycerol, and 0.01% bromphenol blue in 60 mM Tris- HCI, pH 6.8, were denatured a t 100 "C for 5 min. Molecular mass

mately 5 pg of purified wild-type enzyme (WT) and of mutant enzymes standards ( M W ) of the indicated size, expressed in kDa, and approxi-

E558D, E558Q, D766N. and the co-expressed hybrid enzyme E558QI D766N (see text) were subjected to SDS-PAGE using an 8% running gel and a 5% stacking gel. The gel was stained with Coomassie Brilliant Blue R-250.

(Fig. 2). The subunits of wild-type and mutant enzymes had the expected size of approximately 50 kDa (Fig. 2).

Catalysis by Wild-type and Mutant Enzymes-When assayed under conditions optimal for catalysis by the wild-type enzyme, no mutant enzyme exhibited over 0.3% of the activity of wild- type enzyme (Table 11). The activities of mutant enzymes E558D, E558Q, and D766N were also determined at several concentrations of HMG-CoA and NADPH (Fig. 3). Since the K , values of the mutant enzymes for HMG-CoA and NADPH were similar to those for wild-type enzyme (Table 1111, the low V,, values of the mutant enzymes thus cannot be attributed to an inability to bind either substrate. Both G ~ u ~ ~ ~ and Asp766 therefore appear to be catalytically important residues.

In Vitro Complementations-To substantiate the inference that both Glu558 and Asp766 function in catalysis, we asked whether G ~ u ~ ~ ~ and Asp766 mutant enzymes, none of which exhibited over 0.6% of the wild-type activity, might complement one another in vitro. No activity was observed when mutant enzyme E558Q was mixed with mutant enzyme E558D (Table IV). However, upon mixing mutant enzymes E558Q and D766N, or E558D and D766N, activity increased to 1.5 and 6.0% of the wild-type value, respectively (Table IV). The suc-

Active Site of Hamster HMG-CoA Reductase 12 19 TABLE I1

Catalysis by wild-type and mutant enzymes Data are for the reductive deacylation of (SI-HMG-CoA to mevalon-

ate. Oxidation of NADPH was measured spectrophotometrically under conditions optimal for wild-type enzyme, but using up to 0.078 p g of wild-type enzyme, 26 p g of mutant enzymes E558Q and D766N, 29 pg of D766N, and 40 pg of mutant enzyme E558D. Specific activities are expressed both as nanomoles of NADPH oxidized per midmg of protein and as the fraction of the activity of wild-type enzyme.

Enzyme Specific activity

Fraction of wild type activity

Wild-type E558D E558Q D766N

nmol / min / mg 29,200 * 2,800 (100)

%

70 * 1 0.24 85 ? 5 0.29 50 * 8 0.17 $:;p -

0.04 0.02

0 0.04 -0.02 0.0 0.02 0.04 0.06 0.08 -0.4 -0.2 0.0 0.2 0.4 0.6 0.8 1.0

IIWADPH] UpM l/(S)-HMG-CoAl l/W

35 I

30

0 -0.1

I/[NADPH] IIpM

I 6 0 ,

1

-0.1 -0.05 0.0 0.05 0.1 0.15 0.2 0.25 1IWADPHI l/pM

1 0.1

1

30 I I

IO 5

0 -0.54 -0.25 0.0 0.25 0.50 0.75 1.0

: : F l f l , 1 5

0 -0.54 -0.25 0.0 0.25 0.50 0.75 1.0

45 r I

20 15 1501/(/ , , 0 -0.6 -0.3 0.0 0.3 0.6

l/(S)-HhtG-CoA] l/pM

I 0.9

FIG. 3. Dependence of initial velocity on (S)-HMG-CoA and on NADPH concentration. The data are mean values for triplicate de- terminations at pH 6.75 for wild-type enzyme (top) and for mutant enzymes E558Q (middle) and D766N (bottom). Left, assays employed 100 p~ (S)-HMG-CoA (23 times K,) and from 15 to 200 p~ NADPH. Right, assays employed 270 p~ NADPH (8 times K,) and from 1.6 to 16 p~ (SI-HMG-CoA. SA, specific activity, expressed as micromoles of NADPH oxidized per midmg of protein.

TABLE I11 Kinetic parameters for wild-type and mutant enzymes

specific activity (nanomoles of NADPH oxidized per midmg of protein). V,, values, calculated from double-reciprocal plots, are expressed as

Enzyme

nmol lminlmg Wild type 37,200 ? 7,000 4.3 * 0.3 35 * 2 E558D 80 & 20 2.0 * 0.4 11 2 0.5 E5586 78 f 13 D766N

2.3 t 0.3 48 t 14

15 f 2 2.2 2 0.4 17 2 2

W

cessful reconstitution of activity in vitro in the absence of chaotropic agents implies that spontaneous dissociation of the dimeric species into monomers occurred, and that this was followed by subunit exchange.

TABLE IV Catalytic parameters for in vitro reconstituted activities

All data are for the indicated enzyme assayed under conditions optimal for the wild-type enzyme. Specific activities are expressed both as nanomoles of NADPH oxidized per midmg of protein and as the fraction of the activity of wild-type enzyme. Purified wild-type and mutant enzymes were diluted to 3 mg/ml in Buffer A, pH 6.7. Heterodimers were generated by mixing equimolar quantities of each of the mutant enzymes. All samples were incubated on ice overnight prior to assay.

Enzyme Activity

Wild-type nmollminlmg

17,500 % of wild-type

E558D 110 E558Q 80

0.6 0.5

D766N 60 E558D + E558Q 80

0.3

E558D + D766N 1,030 0.5

E558Q + D766N 250 6.0 1.5

(100)

In Vivo Complementation-We next employed a two-plasmid system to co-express two mutant HMG-CoA reductase genes in a single E. coli cell. The expression vectors pACT7 and pT7-7 have the same promoter, but different origins of replication (p15A and ColE1, respectively), and confer different antibiotic resistance phenotypes (&nR and AmpR, respectively) (Fig. 4). To avoid possible recombination during co-expression of two different HMG-CoA reductase genes, E. coli SP1515, a r e d - strain ofE. coli BL21(DE3), was used as the host strain. When expressed individually and purified to electrophoretic homogeneity, mutant enzymes E558Q and D766N exhibited less than 0.6% of wild-type activity (Table VI. However, when both mutant enzymes were co-expressed in the same cell, the resulting purified protein had wild-type affinity for substrates (Table V) and catalyzed HMG-CoAreduction at 25% of the wild-type rate.

Isoelectric Focusing-Wild-type, E558Q, and D766N polypeptides are readily resolved by denaturing isoelectric focussing (Fig. 5). Isoelectric focussing of the co-expressed enzyme revealed the presence of E558Q and D766N subunits, but not of wild-type subunits (Fig. 5). The activity of the co-expressed enzyme thus cannot be attributed to the presence of wild-type enzyme. Rather, the activity of the co-expressed enzyme must reflect the formation of a functional active site from the inactive subunits.

Quaternary Structure-As estimated by size exclusion chromatography, the molecular weight of purified wild-type HMG- CoA reductase is 130,000 2 9,000 (Fig. 6). Based on a molecular weight of 49,618 per subunit, the holoenzyme has 2.6 2 0.2 subunits, a value consistent with a dimer or trimer.

Stoichiometry of HMG-CoA Binding and Determination of the Number of Binding Sites Per Monomer-Approximately 1 mol of HMG-CoA was bound per mol of enzyme monomer (Fig. 7). The Kd of (R,S)-HMG-CoA, 7.6 PM (Fig. 71, matches the K, for (SI-HMG-CoA, 4.3 p~ (Fig. 31, indicating that this K,,, is a true measure of substrate affinity.

DISCUSSION

Based on their strict conservation throughout evolution and on the properties of purified G ~ u ~ ~ ~ and Asp766 mutant enzymes, we infer that acidic residues Glu558 and Asp766 are important for catalysis. Mutagenesis of G ~ u ~ ~ ~ to glutamine and of Asp766 to asparagine resulted in catalytically inactive enzymes. Since the Km values for these mutant enzymes ap- proximated wild-type values, neither G ~ u ~ ~ ~ nor Asp766 appear to function in substrate binding. While a role for Asp766 in catalysis was previously unsuspected, we had anticipated in- volvement of G ~ u ~ ~ ~ since Glus3, the corresponding residue of P mevalonii HMG-CoA reductase, is catalytically significant (3). Future mechanisms proposed for catalysis by HMG-CoA reductase therefore should incorporate roles for 2 acidic residues

1220 Active Site of Hamster HMG-CoA Reductase

Col El pT7-7 (2.5kB)

4 1

AmpR

FIG. 4. Plasmids used for co-expression of mutant enzymes E5586 and D766N. Plasmids pACT7 and pT7-7, which belong to different incompatibility groups, were used to express mutant enzymes D766N and E558Q, respectively. To construct plasmid pACT7, the 920- base pair BamHIIPstI fragment which contains the ampicillin resistance gene of pACYC177 (20) was replaced by the 200-base pair BglIV PstI fragment of pT7-7 (21) which contains the T7 promoter and the pT7-7 polycloning site. The asterisked XbaI site is not unique to either multiple cloning site (MCS) .

TABLE V Kinetic parameters for the heterodimer E558QID766N formed in vivo

All data are for the indicated enzyme assayed under conditions optimal for the wild-type enzyme. K,,, and V,,, values were calculated from plots of l/u versus 1/[S1. Specific activities are expressed both as nanomoles of NADPH oxidized per midmg of protein and as the fraction of the activity of wild-type enzyme.

Enzyme Vmmx Wild-type K,

(S)-HMG-CoA NADPH

% Wild-type 39,000 (100) 4.3 30 E558Q 80 0.2 2.3 14 D766N 50 0.1 2.1 16 E558QD766N 9,600 25 4.7 18

W

Q+N WT Q N QIN +WT WN Q+N

. " I x w -

./ 4

+ FIG. 5. Isoelectric focussing. Isoelectric focussing on a urea-acryl-

amide slab gel was conducted essentially as described by OFarrell(41), but using 8% rather than 4% acrylamide. The gel contained 2% deter- gent (Nonidet P-401, 1.6% pH (X ampholyte, and 0.4% pH 3-10 ampholyte. Focussing was conducted at 200 V for 12 h, then at 800 V for 10 min. Proteins were visualized by staining with Coomassie Brilliant Blue 250. WT, wild-type enzyme; Q. enzyme E558Q; N , enzyme D766N; Q I N , the E558QD766N heterodimer, focussed at two different concentrations; Q+N+WT, a mixture of E558Q. D766N, and wild-type enzymes; Q+N, a mixture of enzymes E5586 and D766N.

rather than 1 as previously suggested (2, 3). That the carboxylate group of both G ~ u ~ ~ ~ and Asp766 per-

form key functions in catalysis was further shown by in vitro and in vivo complementation. We observed significant, but lim- ited, restoration of catalytic activity when inactive mutant enzymes E558Q and D766N were mixed in vitro. Following co- expression in a r e d - strain of E. coli, the purified hybrid enzyme E558Q/D766N exhibited one-quarter of wild-type activity. Since isoelectric focussing confirmed that no wild-type

2.2

2.0 - -

-

\ 2 1.8

$ 1.6

1.4 -

1.2 -

-

1.0 , 0 I I I I I L

0.0 0.5 1.0 1.5 2.0 2.5 3.( log molecular weight

formance liquid size exclusion chromatography. Molecular FIG. 6. Determination of quaternary structure by high per-

weight standards (0) or wild-type enzyme (0) were applied to a 250 x 4.6-mm SynChrom Synchropak GPC size exclusion column and eluted with 200 mM Na,PO,, pH 6.8 (flow rate, 0.5 ml/min), to determine elution volumes, V,. The standards used were thyroglobulin (670 m a ) , bovine y-globulin (158 m a ) , chicken ovalbumin (44 kDa), myoglobin (17 kDa), and vitamin B,* (1.35 m a ) . Blue dextran was used to determine the void volume (V,) and glycyl-tryptophan to determine the total included volume (Vi).

0.14 11

5 0.08

0.021 I I I \ I , 1 o'08.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 B (mole HMG-CoNmole enz. monomer)

FIG. 7. Stoichiometry of HMG-CoA binding to wild-type HMG- CoAreductase. Analyses were conducted in duplicate. The upper com- partment of each ultrafiltration apparatus contained wild-type enzyme (monomer concentration 10.4 p ~ ) and 5.0, 8.0, 35, or 50 p~ (R,S)- [314ClHMG-CoA(specific activity, 440 cpm/pmol) in 1 ml of Buffer A, pH 6.7. Controls lacked either HMG-CoA or enzyme. Following incubation a t 37 "C for 30 min, each apparatus was centrifuged to filter approximately 100 pl. Equal portions of the filtrate, which contains free ligand (F), and the retentate (F plus bound ligand, B ) were then counted. Shown is a plot of ulF versus u, graphed according to the equation ulF = l/Kd(n - u) (42), where u is B divided by the concentration of enzyme monomer (10.4 p i ) . The slope is the reciprocal of the dissociation con- stant, &. The x-intercept gives the number of binding sites per enzyme monomer ( u ) (42).

polypeptide was present, the reconstituted activity can only be due to the E558Q/D766N heterodimer. The high level of in vivo reconstituted activity, 25% that of the wild-type enzyme, indi- cates that half of the co-expressed enzyme consists of E558Q and D766N homodimers, and half of the E558QD766N heterodimer. Only one-half of the active sites in the heterodimer should, however, be active (Fig. 8). Restoration of 25% of wild- type activity thus is the predicted result. The gel filtration data are consistent with a dimeric model, and the stoichiometry of HMG-CoA binding to the wild-type enzyme is consistent with the presence of two active sites per dimer. We therefore con- clude that the functional unit of Syrian hamster HMG-CoA reductase is a dimer with two inter-subunit active sites. In this respect, hamster HMG-CoA reductase resembles ribulose-bi- sphosphate carboxylase (EC 4.1.1.39) (43), mercuric reductase (EC 1.16.1.1) (U), and histidinol dehydrogenase (E.C. 1.1.1.23) (45).

The demonstration that the functionally active form of the catalytic subunit of hamster HMG-CoA reductase is a dimer complements and extends the report of Edwards et al. (12) who

Active Site of Hamster HMG-CoA Reductase 1221

WILD-TYPE ESSSQ

D76hN ES5XQID766N

FIG. 8. Representation of the inter-subunit catalytic site of hamster HMG-CoA reductase. Shown are wild-type enzyme, an inactive homodimer of mutant enzymes E558Q and D766N, and an active heterodimer composed of one E5586 subunit and one D766N subunit. 0 represents a functional active site.

employed radiation-induced inactivation to estimate the size of the membrane-bound rat liver holoenzyme. These investigators concluded that, both under normal conditions and in response to dietary changes that elevate or lower the concentration of HMG-CoA reductase, the membrane-bound rat liver holoenzyme is functionally active as a dimer of noncovalently linked subunits. That the minimum functionally active unit of the membrane-bound mammalian enzyme, and presumably of all eukaryotic HMG-CoA reductases, is dimeric can now be under- stood in light of the present demonstration that both subunits contribute residues essential for catalysis to each site.

Acknowledgments-We thank Robert Simoni of Stanford University for providing a full-length clone of Syrian hamster HMG-CoAreductase; Jon Friesen for carrying out size exclusion chromatography; and Ken- neth Bischoff, Dan Bochar, Jon Friesen, Mark Hermodson, Martin Law- rence, Ramakrishnapillai Omkumar, Scott Rosenthal, and Cynthia Stauffacher for helpful suggestions. We thank Ihor Skrypka and Ronald Somerville for constructing and providing E. coli SP1515, and for very helpful suggestions. Sequence comparisons employed the Wisconsin GCG software resident in the AIDS Center for Computational Biochem- istry at Purdue University, funded by National Institutes of Health Grant AI 127713.

REFERENCES 1. Endo, A. (1988) J. Lipid Res. 33, 1569-1582 2. Veloso. D., Cleland, W. W., and Porter, J. W. (1981) Biochemistry 20,887494 3. Wang, Y., Darnay, B. G., and Rodwell, V. W. (1990) J. Biol. Chem. 266,21634-

4. Darnay, B. G.. Wang, Y., and Rodwell. V. W. (1992) J. Biol. Chem. 267,15064-

5. Mayer, R. J., Debouk, C., and Metcalf, B. W. (1988) Arch. Biochem. Biophys.

6. Ferrer, A,, Aparicio, C., Nogues, N., Wettstein, A, Bach, T. J., and Boronat, A.

21641

15070

267, 110-118

7. Frimpong, K., Darnay, B. G.. and Rodwell, V. W. (1993) Protein Exp. Purif , 4,

8. Darnay, B. G., and Rodwell, V. W. (1993)J. Biol. Chem. 268.8429-8435 9. Ness, G. C., Spindler, C. D., and Momer, M. H. ( 1979) Arch. Biochem. Biophys.

(1990) FEES Letf. 266,67-71

337-344

197.493-499 10. Kleinsek, D. A., and Porter, J. W. (1979) J. Biol. Chem. 254,7591-7599 11. Qureshi, N., Dugan, R. E., Nimmannit, S., Wu. W-H.. and Porter, J. W. (1976)

12. Edwards, P. A,, Kemper, E. S.. Lan, S-F., and Erickson. S. K (1985) J. Biol.

13. Gill, J. F., Jr., Beach, M. J.. and Rodwell, V. W. (1985) J. Biol. Chem. 260,

14. Lawrence. C. M., Rodwell, V. W., and Stauffacher, C. V. (1993) Biophys. J . 64,

15. Bischoff, K M., and Rodwell, V. W. (1992) Biochem. Med. Metab. Biol. 48,

16. Corkey, B. E., Brandt, M., Williams, R. J., and Williamson, J. R. (1981)AnaL

17. Jordan-Starck, T. C., and Rodwell V. W. (1989) J. Bwl. Chem. 264, 17913-

18. Sanger. F., Nicklen, S., and Cou1son.A. R. (1977)Proc. Natl. Acad. Sei. U. S. A.

20. Chang, A. C. Y., and Cohen. S. N. (1978) J. Bacteriol. 134,1141-1156 19. Hsiao, K. (1991) Nucleic Acids Res. 19,2787

21. Studier, F. W.. and Moffatt, B. A. (1986) J. Mol. Biol. 189, 113-130 22. Amersham Corp. (1983) Oligonucleotide-directed in Vitro Mutagenesis System,

Version 2, pp. 1-32, Amersham Corp., Arlington Heights. IL 23. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A

Laboratory Manual, 2nd Ed., Cold Spring Harbor Press, Cold Spring Har- bor, New York

Biochemistry 15,4185-4190

Chern. 260, 10278-10282

9393-9398

A350

149-158

Biochem. 118,3041

17918

74,5463-5467

24. Bradford, M. M. (1976) Anal. Biochem. 72,248-254 25. Promega Corp. (1991) Promega Protocols and Applications Guide, 2nd Ed., pp.

26. Devereux, J., Haeberli. P.. and Smithies, 0. (1984) Nucleic Acids Res. 12, 264-268, Promega Corp., Madison, WI

387-395 27. Chin, D. J., Gil, G., Russell, D. W., Liscum, L., Luskey, K L., Basu, S. K..

Okayama, H., Berg, P., Goldstein, J. L., and Brown, M. S. (1984) Nature 308,613-617

~~ ~~~

28. Skalnik, D. G., and Simoni, R. D. (1985) DNA (N. k:) 4,439-444 29. Luskey, K L.. and Stevens, B. (1985) J. Biol. Chem. 260, 10271-10277 30. Chen. H.. and Shapiro, D. J. (1990) J. B i d . Chem. 265.46224629 31. Woodward. H. D.,Allen, J. M. C., and Lennan, W. J. (1988)J. Biol. Chem. 283,

18411-18418 32. Gertler, F. B.. Chiu, C. Y, Richter-Mann, L., and Chin, D. J. (1988) Mol. Cell.

- . . . - - . . -

Biol. 8. 2713-2721 33. RajkoviqA., Simonsen, J. N., Davis, R. E., and Rottman, F. M. (1989) Proc.

34. Basson, M. E.,Thorsness, M., and Rine, J. (1986)Proc. Natl. Acad. Sci. U. S. A.

35. Basson. M. E.. Thorsness, M., Finer-Moore, J., Stmud, R. M., and Rine. J.

Natl. Acad. Sci. U. S. A. 88,82174221

83,5563-5567

36. Caelles, C., Ferres,A.. Balcells, L., Hegardt, F. G.. and Bor0nat.A. (1989) Plant (1988) Mol. Cell Biol. 8, 37973806

37. Learned, R. M., and Fink, G. R. (1989) Proc. Natl. Acad. Sci. U. S. A. 86,

38. Chye, M., Kush, A,, Tan, C., and Chua, N. (1991) Plant Mol. Biol. 16,567577 39. Lam, W. L., and Doolittle, W. F. (1992) J. Biol. Chem. 267, 5829-5834 40. Beach, M. J., and Rodwell, V. W. (1989) J. Bacteriol. 171,2994-3001 41. OFarrell, P. H. (1975) J. Biol. Chern. 250,40074021 42. Teraoka, H., and Nierhaus, K H. (1979) Methods Enzymol. 69,862466 43. Larimer, F. W.. Lee, E. H., Mural, R. J., Soper, T. S., and Hartman, F. C. (1987)

44. Distefano, M. D., Moore, M. J., and Walsh, C. T (1990) Biochemistry 29,

45. Lee, S. Y., and Grubmeyer, C. T. (1987) J. Bacteriol. 169,3938-3944

Mol. Biol. 13, 627-638

2779-2783

J. Biol. Chem. 262, 15327-15329

2703-2713

the active site of hamster 3-hydroxy-3-methylglutaryl-coa

Documents