Sulfuryl Transfer: The Catalytic Mechanism of Human Estrogen Sulfotransferase*

Huiping ZhangDagger , Olga VarmalovaDagger , Froyland M. VargasDagger , Charles N. Falany§, and Thomas S. LeyhDagger

From the Dagger  Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York 10461 and the § Department of Pharmacology and Toxicology, University of Alabama at Birmingham, Birmingham, Alabama 35294

    ABSTRACT
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Abstract
Introduction
Procedures
Results & Discussion
References

Estrogen sulfotransferase (EST) catalyzes the transfer of the sulfuryl group from 3'-phosphoadenosine 5'-phosphosulfate (PAPS) to 17beta -estradiol (E2). The sulfation of E2 prevents it from binding to, and thereby activating, the estrogen receptor. The regulation of EST appears to be causally linked to tumorigenesis in the breast and endometrium. In this study, recombinant human EST is characterized, and the catalytic mechanism of the transfer reaction is investigated in ligand binding and initial rate experiments. The native enzyme is a dimer of 35-kDa subunits. The apparent equilibrium constant for transfer to E2 is (4.5 ± 0.2) × 103 at pH 6.3 and T = 25 ± 2 °C. Initial rate studies provide the kinetic constants for the reaction and suggest a sequential mechanism. E2 is a partial substrate inhibitor (Ki = 80 ± 5 nM). The binding of two E2 per EST subunit suggests that the partial inhibition occurs through binding at an allosteric site. In addition to providing the dissociation constants for the ligand-enzyme complexes, binding studies demonstrate that each substrate binds independently to the enzyme and that both the E·PAP·E2S and E·PAP·E2 dead-end complexes form. These results strongly suggest a Random Bi Bi mechanism with two dead-end complexes.

    INTRODUCTION
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Abstract
Introduction
Procedures
Results & Discussion
References

Transferring the sulfuryl group (-SO3-) from activated sulfate, or PAPS,1 to a given metabolic recipient typically switches "on" or "off" or otherwise modifies the function of that metabolite (1-6). The extent of sulfation of a given metabolite is determined by the regulated expression of two enzyme classes: the sulfotransferases, which transfer the sulfuryl group, and the sulfatases, which remove it. PAPS appears to be the sole sulfuryl group donor in metabolism. The chemical potential associated with the phosphoric-sulfuric acid anhydride bond of PAPS is remarkably high (Delta Ghydrolysis0' ~ -19 kcal/mol) (7). Thus, the sulfuryl moiety is well positioned energetically for facile and favorable transfer to its metabolic recipients. The chemical and regulatory parallels between sulfation/desulfation and phosphorylation/dephosphorylation are quite strong, yet relatively little is know about the chemistry and enzymology of sulfuryl transfer.

The transcriptional activity of the estrogen receptor (ER) is regulated by sulfation/desulfation at the 3-hydroxyl position of 17beta -estradiol (E2). The binding of E2 to the ER, located in the nuclear membrane, elicits a complex cellular response that is rooted in the transcriptional activity of the ER·E2 complex. E2 binds tightly to the ER (Kd ~ 1 nM) (8, 9); E2S, on the other hand, binds weakly, if at all (10). Thus, the ER-binding activity of estrogen (and, in turn, ER activation) is regulated by E2 sulfation. The sulfation of E2 (Reaction 1),
PAPS<SUP>2−</SUP>+E<SUB>2</SUB>⇌PAP<SUP>2−</SUP>+E<SUB>2</SUB>S<SUP>−</SUP>+H<SUP>+</SUP>
<UP><SC>Reaction</SC> 1</UP>
is catalyzed by the enzyme estrogen sulfotransferase (3'-phosphoadenylylsulfate:estrone 3-sulfotransferase, EC 2.8.2.4). Recent studies indicate that the expression of this enzyme is causally linked to the estrogen-dependent growth response that is believed to underlie the genesis of epithelial breast tumors (11, 12).

Human liver estrogen sulfotransferase (EST) has been cloned and expressed in Escherichia coli K1 (13). In this paper, recombinant EST purified to homogeneity from E. coli is physically characterized. The equilibrium constant for Reaction 1 is determined, and the catalytic mechanism of EST is evaluated in initial rate and ligand binding studies.

    EXPERIMENTAL PROCEDURES
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Materials-- The buffers, salts, enzymes, and reagents, unless specified otherwise, were of the highest grades available from Sigma. [2,4,6,7-3H]Estradiol (85 Ci/mmol) was purchased from NEN Life Science Products. Adenosine 3',5'-[5'-32P]bisphosphate (3000 Ci/mmol) was purchased from ICN Pharmaceuticals. PAPS was purchased from Professor S. Singer (University of Dayton, Dayton, OH). Factor Xa protease was purchased from Enzyme Research Labs. The Bradford assay reagents were purchased from Bio-Rad. The Superdex 200 column was purchased from Amersham Pharmacia Biotech. Amylose resin was obtained from New England Biolabs Inc.

Purification of Estrogen Sulfotransferase-- Competent XL1-Blue (14) cells were transformed with the estrogen sulfotransferase expression vector pMBP-hEST-1 (13) and immediately used to inoculate a 100-ml culture of LB medium (15) containing ampicillin at 50 µg/ml. The pMBP-hEST-1 vector directs the synthesis of a maltose-binding protein-human EST-1 fusion protein, which can be purified by amylose affinity chromatography (13). The culture was incubated overnight at 37 °C and then used to inoculate 10 liters of LB medium to A600 = 0.06. At A600 = 0.6, isopropyl-beta -D-thiogalactopyranoside was added to 350 µM. 3.5 h later, the cells were pelleted at 3500 × g for 25 min, suspended in 500 ml of buffer containing 20 mM Tris-Cl (pH 7.4), 0.20 M KCl, and 5 mM beta -mercaptoethanol, and frozen at -70 °C. All of the purification steps were performed at 4 °C. The cells were thawed, and 1 liter of 4 °C lysis buffer (115 mM Tris-Cl (pH 8.0), 0.375 M sucrose, 0.375 mM EDTA, and 0.03 mg/ml lysozyme) was added. 20 min later, the cells were pelleted at 3500 × g for 25 min. The cell pellet was suspended in sonication buffer (5.0 mM KPO4 (pH 7.4), 1.5 mM DTT, and 57 µM phenylmethylsulfonyl fluoride) and sonicated. The cellular debris was pelleted at 27,000 × g for 60 min, and the concentration of protein, determined by the method of Bradford (16), was adjusted to 3.0 mg/ml using 5.0 mM KPO4 (pH 7.4).

The protein solution was loaded onto a 100-ml bed (3.6 × 17 cm) of amylose resin and equilibrated with 5.0 mM KPO4 (pH 7.4). The column was then washed with 200 ml of wash buffer (10 mM NaH2PO4, 0.20 mM NaCl, 1.0 mM DTT, and 1.0 mM EDTA-HCl (pH 6.3)). The maltose-binding protein-EST fusion protein was then eluted from the column with wash buffer containing 10 mM maltose. 280 A280 units of pure fusion protein was obtained. Glycerol was immediately added to the protein solution to 10% (v/v). Tris-HCl (1.0 M, pH 7.5) was then added to the protein solution to a final concentration of 25 mM. Factor Xa protease (0.66 mg/ml) was added to a solution of maltose-binding protein-EST fusion protein (A280 = 7.0) to a final concentration of 20 µg/ml. The Factor Xa cut site has been engineered into the fusion protein such that the proteolytically produced EST has at its N terminus its initiator methionine (13). The proteolysis was allowed to proceed for 4 h at 4 °C; the reaction reached ~90% completion. The proteolyzed solution was then purified by size-exclusion chromatography using a Superdex 200 (XK 27/70) column equilibrated with 10 mM NaH2PO4 and 1.0 mM DTT. The fractions containing EST were pooled, concentrated using Amicon Centriprep-10 concentrators, and rerun over the Superdex column to remove the remaining traces of fusion protein and maltose-binding protein. The EST was pooled, and the solution was made 10% (v/v) in glycerol. The sample was concentrated to A280 = 1.6. The enzyme was aliquoted, frozen in a dry ice/EtOH bath, and stored at -70 °C. Under these conditions, the enzyme showed no significant loss of activity over the ensuing 3-4 weeks.

Native Molecular Mass-- The native molecular mass of EST was determined by size-exclusion chromatography using a Superdex 200 (XK 27/70) column. The column was equilibrated and run in 50 mM K+-Hepes (pH 8.0) at 4 °C. Bio-Rad gel filtration standards were used to calibrate the column. The apparent native molecular mass of EST was 62 ± 2.3 kDa.

Extinction Coefficient-- The extinction coefficient of EST was determined gravimetrically at 280 nm, the lambda max for the enzyme. EST was dialyzed at 4 °C for 8 h and then overnight against 10 mM NaPO4 (pH 6.3), 1.0 mM KCl, and 0.10 mM DTT. The absorbance of the dialyzed EST was determined in triplicate at 280 nm in a masked 1-cm path length cuvette. Dialyzed enzyme or dialysis buffer (200 ± 1.6 µl) was added to an aluminum weigh boat and dried under vacuum and over P2O5 to a constant weight (<40 h). Triplicate samples of the dialyzed enzyme and buffer were each weighed three times. The absorbance at 280 nm was divided by the concentration of enzyme to calculate the extinction coefficient: epsilon 280* = 1.7 ± 0.1 A280 × (mg/ml)-1 × cm-1.

Equilibrium Constant-- The equilibrium constant for the EST reaction was determined at pH 6.3 and T = 22 ± 2 °C. The measurements were performed in duplicate at each of the following three sets of initial concentrations of E2S, [5'-32P]PAP, and EST, respectively: 100 µM, 240 nM, and 8.0 nM; 200 µM, 240 nM, and 8.0 nM; and 500 µM, 630 nM, and 32 nM. The progress curve for each reaction was determined, and the equilibrium constant was calculated from the reactant concentrations in the stationary phase of the progress curve. The buffer was the same as that used in the initial rate experiments. The concentration of product formed in each reaction was at least 10 times the enzyme active-site concentration. The equilibrium constant was (4.5 ± 0.2) × 103.

Divalent Cation Activation-- The initial rate of the forward reaction was studied as a function of MgCl2 concentration. The assays were performed as described below under "Initial Rate Studies of the Forward Reaction." The conditions of these experiments were as follows: 1.0 nM EST, 10 nM E2, 6.0 µM PAPS, 50 mM KPO4 (pH 6.3), 1.0 mM DTT, and 10% (v/v) glycerol at T = 25 ± 2 °C. The MgCl2 concentration was varied in 2.0 mM increments from 0 to 11 mM. EDTA was added to 2.0 mM in the experiments at zero MgCl2 to ensure that the observed activity was not caused by trace divalent cations in the buffer.

Initial Rate Studies of the Forward Reaction-- The reactions were initiated by adding 50 µl of enzyme to 200 µl of a buffered solution containing varying concentrations of E2 and PAPS. The buffer used in these experiments contained 50 mM KPO4 (pH 6.3), 7.0 mM MgCl2, 1.0 mM DTT, and 10% (v/v) glycerol. The reaction was quenched by the addition of 950 µl of 10 mM KOH. (Controls were run to ensure that the KOH did not hydrolyze the E2S produced.) 5.0 ml of CHCl3 was then added, and the solution was vortexed for 15 s. 800 µl of the aqueous phase was removed and counted. Velocities were determined in duplicate under each of the 16 conditions defined by a 4 × 4 matrix of substrate concentrations. The concentration of EST was 0.10 nM in all of the experiments. The concentrations of PAPS (12, 18, 34, and 300 nM) ranged from 0.2 to 5 times its Km. The concentrations of E2 (2.0, 2.8, 4.8, and 15.5 nM) ranged from 0.4 to 3 times its Km. Partial substrate inhibition by E2 is negligible at these E2 concentrations. The velocities were measured under initial rate conditions since the consumption of the concentration-limiting substrate was <7% of its end point in all cases. The data were statistically fit to the sequen model using the program developed by Cleland (17).

Initial Rate Studies of the Reverse Reaction-- 5.0 µl of EST (4.0 nM) was added to 15 µl of solution containing PAP at varying concentrations and E2S at 250 µM. The reactions were quenched by the addition of 4.0 µl of 0.10 M KOH (final pH 10.0). 9.0 µl of solution was spotted onto polyethyleneimine-fluorescamine TLC plates. The radiolabeled reactants were separated using a LiCl (0.50 M) and HCO2H (2.0 M) mobile phase and quantitated using an AMBIS two-dimensional radioactivity detector. The experiments were performed at 25 ± 2 °C. The data were statistically fit to the hyper model using the weighted least-squares program of Cleland (17).

Substrate Inhibition by E2-- These initial rate assays were performed as described above for the forward reaction. The concentrations of EST and PAPS were 1.0 and 600 nM, respectively. To determine whether inhibition was caused by changes in kcat or the Km for PAPS, the rates at the highest concentrations of E2 (i.e. the plateau region of the curve shown in Fig. 2) were performed at a 10 times higher concentration of PAPS (6.0 µM). Each velocity was determined in duplicate. The data were fit using a partial substrate inhibition model (see "Results and Discussion") with the program Kaleidograph, which uses the Marquardt-Levenberg minimization algorithm.

Fluorescence Titrations-- The EST fluorescence excitation and emission wavelength maxima are 275 and 340 nm, respectively. To avoid possible inner filter effects caused by the absorption of PAP and PAPS, a 285-nm excitation wavelength was used in all of the experiments. The highest nucleotide concentration used in the titrations (5.5 µM) corresponds to an absorbance of 4.7 × 10-4. This absorbance is ~100-fold below the threshold where inner filtering begins to influence intensity measurements (18). The emitted light was detected at 340 nm in all experiments except those involving E2, which is weakly fluorescent when excited at 285 nm. In experiments involving E2, the emission wavelength was set at 360 nm. At this wavelength, the emission from E2 is negligibly low (<0.2% of the EST emission at an equivalent concentration). The solutions used in the titrations were thermally equilibrated and maintained at 25 ± 2 °C during the experiment. The buffer (50 mM KPO4 (pH 6.3), 1.0 mM DTT, 7.0 mM MgCl2, and 10% (v/v) glycerol) was filtered using Gelman 0.2-µm Acrodiscs. The fluorometer used in these studies was a Perkin-Elmer Model LS-5B; the entrance and exit slit widths were set at 10 nm. The titration data were fit to a single-site binding model using the Sigma Plot program, which employs the Marquardt-Levenberg fitting algorithm. The single-site binding model is described by a second order polynomial. The data were fit to the appropriate root of this polynomial to obtain the best fit binding constants.

The protocol of the titration experiments designed to determine Kd differed from those intended to determine stoichiometry. The titrant in the Kd experiments was a solution of EST and ligand in which the ligand concentration was ~10 times the highest ligand concentration reached in the titration. The titrant was added to a concentration-matched solution that did not contain ligand. In the stoichiometry experiments, an EST solution that did not contain ligand was added to a concentration-matched solution that did.

    RESULTS AND DISCUSSION
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Native Molecular Mass and Extinction Coefficient of EST-- The apparent native molecular mass of EST, determined by size-exclusion chromatography (see "Experimental Procedures"), is 62 (±2.3) × 103 Da. The subunit molecular mass, predicted from the DNA sequence of the EST coding region, is 35,123 kDa (13). Thus, the native enzyme appears to be a dimer. The extinction coefficient of EST, determined gravimetrically, is 1.7 ± 0.1 A280 × (mg/ml)-1 × cm-1 (see "Experimental Procedures"). The experimentally determined epsilon 280* is identical to that calculated for EST from its amino acid composition and the epsilon 280* of Trp and Tyr (19).

Stabilizing EST Activity-- In the absence of glycerol, the activity (i.e. the initial rate of PAP and E2S synthesis) of EST is stable for several days at -70 °C, and the half-life of the activity is ~2 h at 25 °C. Glycerol (10%, v/v) prevents detectable deterioration of the activity over 3 h at 25 °C. The addition of E2 to EST in glycerol causes a rapid loss of activity (t1/2 ~ 30 min). This E2-induced inactivation was prevented by the addition of DTT. At 1.0 mM DTT, the activity was not affected at a saturating concentration of E2 over 4 h at 25 °C. Thus, it appears that the binding of E2 potentiates an inactivating oxidation reaction that is suppressed by DTT.

Optimizing Turnover-- Most enzyme-catalyzed transfer reactions involving nucleotides require divalent cations. In these cases, the cations are often directly bound to the polyphosphate chain of the nucleotide. The mechanism of this activation appears to be due predominantly to the entropy reduction associated with the positioning of functional groups for reaction (20). It is interesting that while sulfotransferases catalyze transfer reactions that, in many ways, resemble phosphoryl transfer reactions, they do not require divalent cations for activity. They are, however, activated by divalent cations. A plot of the initial rate of the forward EST reaction versus [MgCl2] is bell-shaped with a maximum at 7.0 mM MgCl2 (see "Experimental Procedures"). The initial rate at zero MgCl2 and 2.0 mM EDTA is 0.18 times the initial rate at 7.0 mM MgCl2. The buffers used in the mechanism studies described in this paper contained MgCl2 at 7.0 mM.

To determine an optimum pH for the EST mechanism studies, the initial rate of the forward reaction was studied as a function of pH at subsaturating E2 and saturating PAPS concentrations. This condition maximizes turnover with respect to both the Km for E2 and kcat. The assay protocol was as described under "Initial Rate Studies of the Forward Reaction", except for the following changes. EST at 0.50 nM and PAPS at 6.0 µM were used, and the pH of the buffer (50 mM KPO4) was varied in 0.4 pH unit increments between 5.4 and 7.4, inclusive, by mixing dibasic and monobasic solutions of KPO4. The pH rate profile was bell-shaped with a maximum initial rate at pH 6.3. The recent structure of mouse testis estrogen sulfotransferase implicates His-108 as a general base that abstracts a proton from the 3-hydroxyl group of E2, thereby activating it for attack (21). Further pH rate studies will help to determine whether this residue contributes to the pH rate dependence of the EST-catalyzed reaction.

Equilibrium Constant-- To evaluate the energetics associated with transferring the sulfuryl group between PAPS and E2 and to aid in the design of initial rate experiments, the equilibrium constant for the EST reaction was determined at pH 6.3 and T = 25 ± 2 °C (the conditions of the initial rate studies). The equilibrium constants were calculated from reactant concentrations in the stationary phases of reaction progress curves constructed in duplicate at three different sets of E2S and PAP concentrations (see "Experimental Procedures"). Controls were run to ensure that the EST activity did not change during the experiments. The equilibrium constant is (4.5 ± 0.2) × 103. The Delta G0 associated with this Keq is -5.0 kcal/mol. It should be emphasized this apparent equilibrium constant does not explicitly include the proton and divalent cation dependences.

Initial Rate Study of the Forward Reaction-- To determine the kinetic constants for the forward reaction and to obtain a preliminary assessment of the order of substrate binding, a classical initial rate study of the forward reaction was performed. The initial rate was determined as a function of both E2 and PAPS concentrations (see "Experimental Procedures"). The results of this study are shown in Fig. 1. The pattern of the data is indicative of a sequential mechanism (one in which both substrates must bind to the enzyme before product is released). However, it does not rule out a ping-pong mechanism with an unstable enzyme intermediate. An equilibrium ordered mechanism is ruled out by the fact that the lines through the data of the 1/V versus 1/[E2] and 1/V versus 1/[PAPS] (not shown) plots do not intersect on the 1/V axis (22). The kinetic constants obtained from this study are compiled in Table I. kcat (1.3 ± 0.08 s-1) and the Km for PAPS (59 ± 13 nM) are similar to those measured for other sulfotransferases (23, 24). The Km for E2 is comparable to the in vivo concentration of E2, ~1 nM (25), suggesting that the enzyme is optimized to perform at the physiological concentration of E2.


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Fig. 1.   An initial rate study of the synthesis of PAP and E2. The initial rate of PAP and E2S synthesis is shown as a function of PAPS and E2 concentrations. The PAPS concentration was varied between 0.20 and 5.0 times its Km (59 nM). The E2 concentration was varied between 0.4 and 3 times its Km (5.2 nM). The E2 concentration range was sufficiently low that inhibition by E2 was negligible. Each point represents the average of two independent determinations. The lines through the points represent the best fits to the data. The experiments were performed at 25 ± 2 °C. For the experimental protocol, see "Experimental Procedures."

                              
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Table I
Kinetic constants

Initial Rate Study of the Reverse Reaction-- The Km for PAP and kcat for the reverse reaction were determined in an initial rate study at a saturating (920 × Kd) concentration of E2S (250 µM). Controls were run to ensure that E2S did not inhibit the velocity at this concentration. The Km for PAP was 38 ± 0.8 nM, and kcat was 0.16 ± 0.0013 min-1 (Table I). The experimental protocol is described under "Experimental Procedures." Given the technical obstacles associated with the unfavorable equilibrium constant for the reverse reaction and the relatively high Km for E2S, the order of substrate addition for the reverse reaction was determined with the equilibrium binding studies described below.

Partial Substrate Inhibition by E2-- The initial rate data shown in Fig. 2 demonstrate that E2 inhibits the forward reaction. The fact that the velocity decreases to a plateau, rather than to zero, means that one or more of the kinetic parameters for the reaction are being titrated from one value to another as E2 adds to the enzyme. The inhibition experiment (Fig. 2, bullet ) was performed at a fixed near-saturating concentration of PAPS (600 nM, 10 × Kd). If the inhibition were due solely to an increase in the Km for PAPS, causing the concentration of the reactive form(s) of the enzyme to decrease, increasing the concentration of PAPS would drive the initial rates, at inhibitory concentrations of E2, back to the uninhibited levels. This, in fact, does not occur. Increasing the concentration of PAPS 10-fold (Fig. 2, square ) had no significant effect on the initial rates at high E2 concentrations. Thus, it is kcat that is affected by the binding of E2 in these experiments.


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Fig. 2.   Partial substrate inhibition by E2. The initial rate of E2S and PAP synthesis was determined as a function of E2 concentration. The experimental conditions are described under "Experimental Procedures." bullet , data obtained at EST and PAPS concentrations of 1.0 and 600 nM, respectively; square , velocities determined at 6.0 µM PAPS. Each velocity is the average of at least two independent determinations. The solid line passing through the points represents the initial rate profile predicted by the best fit model.

The Ki for E2 was evaluated by fitting the data shown in Fig. 2 to the algebra that describes the kinetic behavior of the model shown in Fig. 3. In the model, all of the enzyme forms are saturated with PAPS. The kinetic constants V1 and Km for E2 were obtained from the initial rate study of the forward reaction (Table I). V2 was set at 0.18 nM/min, which is slightly below the plateau shown in Fig. 2. The data were fit to the following equation: v = V1(1 + (V2[E2]/V1Ki))/(1 + Km/[E2] + [E2]/Ki) (26). This equation assumes that substrate binding is at equilibrium, which is plausible given the very low turnover of the enzyme. The value of Ki that provides the best fit to the data is 80 ± 5 nM. It should be mentioned that the inhibition model was used to select the E2 concentrations used in the initial rate studies (Fig. 1) such that inhibition by E2 was negligible.


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Fig. 3.   Kinetic model for partial substrate inhibition by E2. Km was obtained from Table I; V1 was calculated from kcat (Table I), and V2 was set at 0.18 min-1, slightly below the initial rate in the plateau of Fig. 2. The algebra that describes the kinetic behavior of this model was used to obtain a best fit value for Ki.

Equilibrium Binding Studies-- The excitation and emission wavelength maxima for EST are 275 and 340 nm, respectively. The fine structure and lambda max of the emission spectrum do not change significantly when substrates bind to EST; however, the intensity decreases 30-50% depending on the ligand. These ligand-dependent decreases in the quantum yield of EST provide excellent experimental handles to determine both the equilibrium constants and stoichiometry of the enzyme-ligand interactions.

The stoichiometry of the enzyme-ligand interactions was determined in fluorescence titration experiments in which the concentration of enzyme active sites was >15 × Kd. At these enzyme concentrations, the binding isotherms have two linear regions. They are linear in the substrate concentration range 0~0.4 × [enzyme] because virtually all of the substrate is bound to the enzyme in this range, and they are linear (with a slope of zero) at saturating substrate concentrations. The intersection of the lines extrapolated from these linear regions corresponds, on the [ligand]/[E] axis, to the stoichiometry of the ligand-enzyme complex. The results of these studies for each of the EST substrates are depicted in Fig. 4 (A-D). The stoichiometries of the ligand-enzyme complexes are 1:1 for all of the ligands except E2, which has a stoichiometry of 2:1. The binding of two E2 to each EST subunit strongly suggests that one of the binding sites is the catalytic site, whereas the other is the allosteric site that regulates the turnover of the enzyme (Fig. 3). Notwithstanding the possibility that these fluorescence experiments monitor the formation of nonproductive complexes, the independent binding of each of the EST substrates demonstrates that the mechanism of the enzyme is random sequential.


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Fig. 4.   Stoichiometry of ligand binding. The stoichiometries were determined by monitoring the change in fluorescent intensity of the enzyme as a function of ligand concentration at enzyme active-site concentrations >14 × Kd. I/I0 is the ratio of the fluorescence intensity of the enzyme at a given ligand concentration to that in the absence of ligand. The straight lines through the points were fit by eye. Each point represents the average of at least two independent determinations. The PAP, PAPS, E2S, and E2 titrations are shown in A-D, respectively. The EST concentrations used in the experiments associated with A-D were 1.5, 2.0, 2.0, and 1.0 µM, respectively.

The equilibrium constants for the formation of enzyme-substrate complexes were determined by fluorescence titrations in which the enzyme concentration was held fixed between 0.4 and 5 × Kd. The binary complex binding constants were determined by fitting the data shown in Fig. 5 (A, C, and D) to a single-site binding model. The Kd values for the E·PAPS, E·PAP, and E·E2S complexes are 37 ± 0.3, 30 ± 0.3, and 271 ± 49 nM, respectively (Table II). The data associated with the binding of E2 (Fig. 5B) are equally well fit using either a single-site model (Kd = 26 ± 2 nM; shown in Fig. 5B) or a two-site model that assumes no interaction energy (27) in which the dissociation constants are within an order of magnitude of one another and symmetrically disposed about the best fit Kd predicted by the single-site model.


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Fig. 5.   Binary and ternary (dead-end) complex formation. The affinities of enzyme-ligand interactions were determined by monitoring changes in the intrinsic fluorescence of EST as a function of ligand concentration. I/I0 is the ratio of the fluorescence intensity of the enzyme at a given ligand concentration to that in the absence of ligand. The curves through the points represent the binding isotherms predicted by the best fit parameters obtained by fitting the experimental data to a single-site binding model. Each point represents the average of two to three independently determined values. The binary complexes were as follows: A, PAPS binding to EST; B, E2 binding to EST; C, PAP binding to EST; D, E2S binding to EST. The ternary (dead-end) complexes were as follows: E, PAPS binding to EST·E2S; F, PAP binding to EST·E2. The EST concentrations associated with A-F were 75, 100, 75, 117, 150, and 100 nM, respectively. The concentrations of E2S and E2 used in the ternary complex experiments were 6.0 µM (22 × Kd) and 1.5 µM (>30 × Kd), respectively.

                              
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Table II
Dissociation constants

The formation of the ternary dead-end complexes was also investigated using fluorescence titrations (Fig. 5, E and F). The binding of PAPS to the E·E2S complex and of PAP to the E·E2 complex was monitored at a saturating concentration of E2S or E2. The data clearly demonstrate that both dead-end complexes form. The nucleotide dissociation constants for the E·PAPS·E2S and E·PAP·E2 complexes are 20 ± 2.8 and 22 ± 1.7 nM, respectively. Comparison of the binary and dead-end dissociation constants for PAPS and PAP reveals a slight binding synergism between the ligands in the ternary complex (Table II). These results corroborate the dead-end complexes implicated by earlier product inhibition studies with arylsulfotransferase (24) and strongly suggest that the mechanism of EST is Random Bi Bi with two dead-end complexes.

Conclusions-- Initial rate and ligand binding experiments have been used to investigate the catalytic mechanism of EST. The kinetic parameters for the mechanism were determined from the initial rate studies, which also suggested that the mechanism is sequential. Ligand binding studies were used to determine the equilibrium constants and stoichiometries of the enzyme-substrate interactions. The binding studies demonstrated that each of the substrates can bind independently to the enzyme and that two dead-end complexes can form. These results strongly suggest a Random Bi Bi mechanism with two dead-end complexes. The initial rate experiments revealed that E2 is a partial substrate inhibitor of the reaction with a Ki of 80 ± 5 nM. The mechanism of the inhibition is partially delineated by the stoichiometry studies, which show that the enzyme contains two E2-binding sites/catalytic subunit, suggesting that the enzyme harbors an allosteric E2-binding site.

    FOOTNOTES

* This work was supported by National Institutes of Health Grants GM54469 (to T. S. L.) and GM38953 (to C. N. F.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Dept. of Biochemistry, Albert Einstein College of Medicine, 1300 Morris Park Ave., Bronx, NY 10461. Tel.: 718-430-2857; Fax: 718-430-8565.

1 The abbreviations used are: PAPS, 3'-phosphoadenosine 5'-phosphosulfate; PAP, adenosine 3',5'-diphosphate; ER, estrogen receptor; E2, 17beta -estradiol; E2S, 17beta -estradiol 3-sulfate; EST, estrogen sulfotransferase; DTT, dithiothreitol.

    REFERENCES
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Results & Discussion
References

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