3'-Phosphoadenosine 5'-phosphosulfate allosterically regulates sulfotransferase turnover

Abstract

Human cytosolic sulfotransferases (SULTs) regulate the activities of thousands of small molecules-metabolites, drugs, and other xenobiotics-via the transfer of the sulfuryl moiety (-SO3) from 3'-phosphoadenosine 5'-phosphosulfate (PAPS) to the hydroxyls and primary amines of acceptors. SULT1A1 is the most abundant SULT in liver and has the broadest substrate spectrum of any SULT. Here we present the discovery of a new form of SULT1A1 allosteric regulation that modulates the catalytic efficiency of the enzyme over a 130-fold dynamic range. The molecular basis of the regulation is explored in detail and is shown to be rooted in an energetic coupling between the active-site caps of adjacent subunits in the SULT1A1 dimer. The first nucleotide to bind causes closure of the cap to which it is bound and at the same time stabilizes the cap in the adjacent subunit in the open position. Binding of the second nucleotide causes both caps to open. Cap closure sterically controls active-site access of the nucleotide and acceptor; consequently, the structural changes in the cap that occur as a function of nucleotide occupancy lead to changes in the substrate affinities and turnover of the enzyme. PAPS levels in tissues from a variety of organs suggest that the catalytic efficiency of the enzyme varies across tissues over the full 130-fold range and that efficiency is greatest in those tissues that experience the greatest xenobiotic "load".

Figure 1

Equilibrium binding of PAPS to SULT1A1. (A) PAPS binding to the high-affinity subunit. Binding was monitored via ligand-induced changes in the intrinsic fluorescence of SULT1A1 (λ_ex = 295 nm; λ_em = 345 nm). Reaction conditions included SULT1A1 (0.05 μM, dimer), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2 °C. Each point is the average of three independent determinations. The solid line through the data represents a least-squares fit using a model that assumes a single binding site per dimer. K_d = 0.37 ± 0.05 μM. (B) PAPS binding stoichiometry at the high-affinity site. The conditions were identical to those in described for panel A except that [SULT1A1] = 3.0 μM dimer (16K_d). The stoichiometry was 1.1 ± 0.2 PAPS molecules per dimer. (C) PAPS binding at the low-affinity site. Experimental conditions were identical to those in described for panel B. PAPS binding is biphasic. The high- and low-affinity phases are colored red (inset) and black, respectively. The line through the points represents a least-squares fit to the low-affinity phase using a model that assumes a single binding site per dimer. K_d = 30 ± 4 μM. (D) Full-site PAPS binding stoichiometry. The reaction conditions were identical to those described for panel A except that [SULT1A1] = 475 μM dimer (16K_d for the low-affinity site). The stoichiometry was 2.1 ± 0.2 PAPS molecules per dimer, or 1.1 ± 0.1 per subunit.

Figure 2

Pre-steady-state binding of PAPS to SULT1A1. (A) Composite k_obs vs [PAPS] plot. Two well-isolated binding phases are observed. Binding was monitored via changes in SULT1A1 intrinsic fluorescence (λ_ex = 290 nm; λ_em ≥ 330 nm). k_obs values are the average of three independent determinations. Reaction conditions included SULT1A1 (0.050 μM, dimer), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2 °C. Red dots indicate the k_obs values predicted using the k_on and k_off values obtained from the experiments associated with panels B and C. (B) k_obs vs [PAPS] for the high-affinity subunit. Reaction conditions were identical to those described for panel A except that [SULT1A1] = 0.030 μM (dimer). k_on = 2.0 ± 0.2 μM^–1 s^–1; k_off = 0.70 ± 0.02 s^–1. (C) k_obs vs [PAPS] for the low-affinity subunit. Reaction conditions were identical to those described for panel A except the SULT1A1 (2.0 μM, dimer) was equilibrated with PAPS [8.0 μM, 26K_{d(high affinity)}, 0.27K_{d(low affinity)}] before being mixed with PAPS at higher concentrations (20–80 μM). k_on = 0.96 ± 0.01 μM s^–1; k_off = 29 ± 1 s^–1. All reactions were pseudo-first-order in PAPS concentration.

Figure 3

Binding of TAM to E and E·(PAP)₂. (A) TAM binding to E. Binding was monitored via changes in SULT1A1 intrinsic fluorescence (λ_ex = 290 nm; λ_em = 345 nm). Reaction conditions included SULT1A1 (0.10 μM, dimer), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2 °C. Each point is the average of three independent determinations. The curve is the behavior predicted by a best fit model that assumes a single binding site per dimer. K_d = 0.67 ± 0.04. (B) TAM binding to E(PAP)₂. Conditions and data analysis were identical to those described for panel A except PAP = 0.50 mM (17K_d for PAPS binding at its low-affinity site). The K_d for TAM binding is 0.68 ± 0.12 μM. (C) Stoichiometry of binding of TAM to E and E·(PAP)₂. Conditions were identical to those described for panels A and B except that [SULT1A1] = 10 μM (dimer). Binding to E and E·(PAP)₂ is shown with filled and empty circles, respectively. The stoichiometries are 2.0 ± 0.1 TAM bound per SULT1A1 dimer.

Figure 4

Binding of TAM to SULT1A1. (A) TAM binding is biphasic. Binding was monitored via changes in SULT1A1 intrinsic fluorescence (λ_ex = 290 nm; λ_em = 345 nm). The titration conditions included SULT1A1 (0.10 μM, dimer), PAP (6.0 μM), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2 °C. The distribution of enzyme forms at 6.0 μM PAP is as follows: E·PAP, 79%; E·(PAP)₂, 16%; E, 5.0%. The first phase (red dots, inset) shows binding of TAM to the PAP-free subunit of SULT1A1 (K_d = 0.67 ± 0.05 μM). The second phase shows binding of TAM to the nucleotide-bound subunit (K_d = 13 ± 2 μM). Each point is the average of two independent determinations. The line through the points is the behavior predicted by a best-fit model that assumes a single binding site per dimer. The first and second phases were fit separately using the data indicated by the red and black circles, respectively. (B) Semiquantitative stoichiometric binding of TAM. The conditions of the titration included SULT1A1 (9.0 μM, dimer), PAP (9.0 μM), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2 °C. At these concentrations, the distribution of enzyme forms is as follows: E·PAP, 77%; E·(PAP)₂, 3.8%; E, 19%. The binding is biphasic. The first phase (red dots, inset) indicates a stoichiometry of approximately one TAM binding site per dimer. A second low-affinity phase is also observed (black dots).

Figure 5

SUTL1A1 turnover as a function of PAPS occupancy. A plot of SULT1A turnover vs [PAPS] is biphasic. The first and second phases correspond to saturation of the high- and low-affinity PAPS binding sites, respectively. The reaction was monitored via 1-HPS fluorescence (λ_ex = 320 nm; λ_em = 380 nm). The conditions included SULT1A1 (1.0 nM, dimer), 1-HP (160 μM, 20K_d), MgCl₂ (5.0 mM), NaPO₄ (50 mM), pH 7.2, and 25 ± 2.0 °C. The asterisks indicate the PAPS concentrations (0.20 and 2.0 μM) used in initial rate studies to obtain Michaelis parameters for the E·PAP and E·(PAPS)₂ forms.

Figure 6

Predicted fraction of E·(PAPS)₂ in human tissues. Fractions were calculated using reported PAPS concentrations.⁻

Figure 7

Coupling of PAPS binding and cap closure in SULT1A1. The ligand binding sites of the unliganded enzyme are open and can receive ligands. Binding of the first PAPS molecule closes both the PAPS and acceptor binding sites of the subunit to which PAPS has bound. In this configuration, PAPS cannot escape and only small acceptors can enter unless the enzyme isomerizes to the open form (not shown), which is unfavorable (K_iso = 26 in favor of the closed state). Consequently, the singly PAPS-bound configuration favors small acceptors. As the second PAPS molecule binds, all of the binding sites open, thus alleviating the catalytic bias against large substrates, and each subunit turns over 4-fold faster.