The bicarbonate/carbon dioxide pair increases hydrogen peroxide-mediated hyperoxidation of human peroxiredoxin 1

Abstract

2-Cys peroxiredoxins (Prxs) rapidly reduce H₂O₂, thereby acting as antioxidants and also as sensors and transmitters of H₂O₂ signals in cells. Interestingly, eukaryotic 2-Cys Prxs lose their peroxidase activity at high H₂O₂ levels. Under these conditions, H₂O₂ oxidizes the sulfenic acid derivative of the Prx peroxidatic Cys (C_PSOH) to the sulfinate (C_PSO₂^-) and sulfonated (C_PSO₃^-) forms, redirecting the C_PSOH intermediate from the catalytic cycle to the hyperoxidation/inactivation pathway. The susceptibility of 2-Cys Prxs to hyperoxidation varies greatly and depends on structural features that affect the lifetime of the C_PSOH intermediate. Among the human Prxs, Prx1 has an intermediate susceptibility to H₂O₂ and was selected here to investigate the effect of a physiological concentration of HCO₃^-/CO₂ (25 mm) on its hyperoxidation. Immunoblotting and kinetic and MS/MS experiments revealed that HCO₃^-/CO₂ increases Prx1 hyperoxidation and inactivation both in the presence of excess H₂O₂ and during enzymatic (NADPH/thioredoxin reductase/thioredoxin) and chemical (DTT) turnover. We hypothesized that the stimulating effect of HCO₃^-/CO₂ was due to HCO₄^-, a peroxide present in equilibrated solutions of H₂O₂ and HCO₃^-/CO₂ Indeed, additional experiments and calculations uncovered that HCO₄^- oxidizes C_PSOH to C_PSO₂^- with a second-order rate constant 2 orders of magnitude higher than that of H₂O₂ ((1.5 ± 0.1) × 10⁵ and (2.9 ± 0.2) × 10³ m^-1·s^-1, respectively) and that HCO₄^- is 250 times more efficient than H₂O₂ at inactivating 1% Prx1 per turnover. The fact that the biologically ubiquitous HCO₃^-/CO₂ pair stimulates Prx1 hyperoxidation and inactivation bears relevance to Prx1 functions beyond its antioxidant activity.

Keywords: antioxidant defense; bicarbonate; carbon dioxide; hydrogen peroxide; hyperoxidation; inactivation; peroxiredoxin; peroxymonocarbonate; redox signaling; thiol; thiol oxidation.

Figure 1.

Catalytic cycle and the hyperoxidation pathway of 2-Cys peroxiredoxins (A) and effects of HCO₃⁻/CO₂ on Prx1 hyperoxidation (B). A, 2-Cys Prx is represented by its minimal functional unit, which is a homodimer, with one of the peroxidatic (C_PS⁻) and one of the resolving Cys (C_RSH) shown. The catalytic cycle starts with the rapid reaction of the peroxidatic cysteine with hydroperoxides to reduce them, while being oxidized to the sulfenic acid form (C_PSOH). Next, the resolving Cys reacts with C_PSOH of the adjacent monomer forming a head–to–tail disulfide (C_PS–SC_R). This oxidized form of Prx is reduced by the NADPH/Trx/TrxR system, completing the peroxidase cycle. Because a partial unfolding of the α-helix containing C_P is required for the formation of the disulfide, C_PSOH can be further oxidized to the sulfinate (C_PSO₂⁻) and, subsequently, to the sulfonated (C_PSO₃⁻) forms if the oxidant concentration is sufficiently high. These processes are called hyperoxidation and lead to inactivation of the peroxidase activity of the enzyme. B, representative effects of HCO₃⁻/CO₂ on H₂O₂-mediated Prx1 hyperoxidation revealed by nonreducing SDS-PAGE (lower panel) and Western blot analysis using an anti-PrxSO₂H/PrxSO₃H antibody (upper panel). The incubations contained Prx1 (2.5 μm), H₂O₂ (2.5–100 μm) in the absence and presence of HCO₃⁻/CO₂ (25 mm) in phosphate buffer (50 mm) containing DTPA (0.1 mm), pH 7.4. After a 5-min incubation at room temperature, the samples were treated and analyzed as described under “Experimental procedures.” The shown experiment is representative of three independent experiments.

Figure 2.

Stimulatory effect of HCO₃⁻/CO₂ (25 mm) on H₂O₂-mediated Prx1 hyperoxidation during enzymatic turnover. A, representative kinetics of NADPH consumption coupled to the reduction of the specified concentrations of H₂O₂ by Prx1 (0.6 μm) turnover by NADPH (200 μm), Trx1 (2.5 μm), TrxR1 (80 nm) in Hepes-NaOH (50 mm) containing 1 mm EDTA, pH 7.0, at 30 °C. B, same as A in the presence of HCO₃⁻/CO₂ (25 mm). C, comparison of the kinetics of NADPH consumption coupled to the reduction of H₂O₂ (0.5 mm) in the absence (black line) or presence of HCO₃⁻/CO₂ (25 mm) (red line). Experimental conditions are the same as A and B. D, plots of f_inactivation versus H₂O₂ concentration in the absence and presence of HCO₃⁻/CO₂ (25 mm) yielded straight lines, and the slope was used to calculate C_{1%hyperoxidation} for H₂O₂ in the absence ((335 ± 18) μm) and presence of HCO₃⁻/CO₂ (25 mm) ((101 ± 4) μm). The values of f_inactivation were obtained by treatment of the kinetic data obtained in experiments similar to A and B as described previously (8) and under “Experimental procedures”; the values plotted in D are the mean ± S.D. obtained from three independent kinetic experiments.

Figure 3.

Effect of H₂O₂, HCO₃⁻/CO₂, or H₂O₂ plus HCO₃⁻/CO₂ (25 mm) on the insulin reductase activity of NADPH/TrxR/Trx (A) and on Prx1 hyperoxidation during enzymatic turnover (B). A, kinetics of NADPH consumption and insulin precipitation during the insulin reductase activity of the TrxR/Trx system. The incubation contained NADPH (200 μm), Trx1 (2.5 μm), TrxR1 (80 nm), and insulin (0.16 mm) in Hepes-NaOH (50 mm) containing 1 mm EDTA, pH 7.0, at 30 °C (black curve); the same plus H₂O₂ (0.5 mm) (blue curve); the same plus HCO₃⁻/CO₂ (25 mm) (gray curve); and the same plus H₂O₂ (0.5 mm) and HCO₃⁻/CO₂ (25 mm) (red curve). The absorbance values shown are the mean ± S.D. obtained from three independent experiments. B, representative nonreducing SDS-PAGE (left panel) and Western blot analysis using an anti-PrxSO₂H/PrxSO₃H antibody (right panel) of Prx1 hyperoxidation during enzymatic turnover. The incubations contained Prx1 (0.6 μm), NADPH (200 μm), Trx1 (2.5 μm), TrxR1 (80 nm), and the specified concentrations of H₂O₂ in Hepes-NaOH (50 mm) containing 1 mm EDTA in the absence or presence of HCO₃⁻/CO₂ (25 mm), pH 7.0; the controls with HCO₃⁻/CO₂ (25 mm) are also shown. After 15 min at 30 °C, the samples were treated and analyzed as described under “Experimental procedures.” The shown experiment is representative of several similar experiments, including some performed with an antibody from a different commercial source (Fig. S1).

Figure 4.

Nano-ESI–Q-TOF–MS/MS analysis of the alkylated, sulfinated, and sulfonated peptides ³⁸YVVFFFYPLDFTFVCPTEIIAFSDR⁶² from tryptic digests of Prx1 after chemical turnover. Prx1 (5 μm) was treated with 1 mm H₂O₂ plus HCO₃⁻/CO₂ (25 mm) in the presence of DTT (10 mm) for 16 h at 30 °C. The incubations were performed in phosphate buffer (50 mm) containing DTPA (0.1 mm), pH 7.4, and the samples were treated and analyzed as described under “Experimental procedures.” A, MS/MS sequencing of the peak at m/z value of 1054.5171, which corresponds to the alkylated peptide (monoisotopic mass 3160.5290) with three charges, found in the tryptic digests of treated Prx1. R represents the NEM (–C₆H₈NO₂) group. B, MS/MS sequencing of the peak at m/z value of 1023.4979, which corresponds to the sulfinated (–SO₂H) peptide (monoisotopic mass 3067.4711) with three charges, found in the tryptic digests of treated Prx1. R represents the –SO₂H group. C, MS/MS sequencing of the peak at m/z value of 1028.8295, which corresponds to the sulfonated (–SO₃H) peptide (monoisotopic mass 3083.4661) with three charges, found in the tryptic digests of treated Prx1. R represents the –SO₃H group. The spectra shown are representative of three independent experiments.

Figure 5.

Relative yields of alkylated, sulfinated, and sulfonated peptides obtained from tryptic digests of Prx1 after chemical turnover. Prx1 (5 μm) was untreated (black bars) or treated with 1 mm H₂O₂ (blue bars) or with 1 mm H₂O₂ plus HCO₃⁻/CO₂ (25 mm) (red bars) in the presence of DTT (10 mm) for 16 h at 30 °C. The incubations were performed in phosphate buffer (50 mm) containing DTPA (0.1 mm), pH 7.4, and the samples were treated and analyzed as described under “Experimental procedures.” A, sum of the specified peptides containing Cys⁵² (C_P). B, sum of the specified peptides containing Cys¹⁷³ (C_R). C, sum of the specified peptides containing Cys⁷¹ and Cys⁸³. The values shown are the mean ± S.D. obtained from three independent experiments; the calculations were performed as described in the text.

Figure 6.

Evaluating the role of HCO₄⁻ in the stimulatory effect of HCO₃⁻/CO₂ on H₂O₂-mediated Prx1 hyperoxidation. A, determination of C_{1% hyperoxidation} for HCO₄⁻ based on the kinetic data of Fig. 2D and the calculations described in the text. As in Fig. 2D, the f_inactivation values plotted are the mean ± S.D. obtained from three independent experiments. B, representative increase in the intrinsic fluorescence of oxidized Prx1 (Prx1 C_PSOH) over time following the addition of 25 μm (black trace), 1 mm (red trace), and 2 mm (blue trace) H₂O₂; the initial concentration of reduced Prx1 was 2.5 μm. The inset shows the changes in the intrinsic fluorescence of reduced Prx1 (2.5 μm) over time following the addition of 2.5 μm H₂O₂. C, determination of the second-order rate constant of the reaction between Prx1 C_PSOH and H₂O₂ to produce Prx1 C_PSO₂⁻. The values of k_obs were obtained by fitting the traces, similar to those shown in B, to a single exponential equation. Plotting the values of k_obs versus H₂O₂ concentration yielded a straight line, with the slope providing the apparent second-order rate constant of 2.9 ± 0.2 × 10³ m⁻¹ ·s⁻¹. The k_obs values plotted in C are the mean ± S.D. obtained from three independent experiments. D, determination of the second-order rate constant of the reaction between Prx1 C_PSOH and HCO₄⁻ to produce Prx1 C_PSO₂⁻. The values of k_obs were obtained by fitting the traces obtained in experiments similar to those shown in B but with H₂O₂ (0.5 mm) and variable concentrations of HCO₃⁻/CO₂ (25–100 mm) to a single exponential equation. Plotting the values of k_obs against HCO₄⁻ concentrations calculated from Equation 1 yielded a straight line, the slop of which provided the apparent second-order rate constant of (1.5 ± 0.1) × 10⁵ m⁻¹·s⁻¹. The k_obs values plotted in D are the mean ± S.D. obtained from three independent experiments. The reactions in B–D were performed in phosphate buffer (50 mm) containing 0.1 mm DTPA, pH 7.4, at 25 °C as described under “Experimental procedures.”

Figure 7.

Schematic representation of the equilibria involved in HCO₄⁻ formation from H₂O₂ and HCO₃⁻/CO₂ (A) and of the mechanism proposed for the stimulatory effect of HCO₃⁻/CO₂ on H₂O₂-mediated Prx1 hyperoxidation and inactivation is shown. A, mechanism of HCO₄⁻ formation from H₂O₂/HOO⁻ was from Richardson and co-workers (43); the rate constants of all proposed reactions (Table S1) were used to perform the computer simulation HCO₄⁻ formation from H₂O₂ (0.5 mm) and HCO₃⁻/CO₂ (25 mm) displayed in Fig. S4. B, stimulatory effect of HCO₃⁻/CO₂ on H₂O₂-mediated Prx1 hyperoxidation is attributed to HCO₄⁻. 2-Cys Prx is represented by its minimal functional unit, which is a homodimer, with one of the peroxidatic (C_PS⁻) and one of the resolving Cys (C_RSH) shown. HCO₄⁻ coexists in H₂O₂ solutions containing HCO₃⁻/CO₂ (Fig. 6A) and oxidizes C_PSOH to C_PSO₂⁻ with a second-order rate constant 2 orders of magnitude higher than that of H₂O₂. HCO₄⁻ should not influence the first step of the Prx1 catalytic cycle, because the reduction of H₂O₂ and concomitant oxidation of C_PS⁻ to C_PSOH is extremely rapid. Prx1 hyperoxidation is an inactivation pathway that depends on specific Prx1 structural features that determine the lifetime of the C_PSOH intermediate. Because HCO₄⁻ is considerably more reactive toward the C_PSOH intermediate than H₂O₂, it will redirect higher levels of C_PSOH from the catalytic cycle to the hyperoxidation and inactivation pathway. Our data suggested but did not prove that HCO₄⁻ oxidizes C_PSO₂H to C_PSO₃H more rapidly than H₂O₂ justifying the use of broken lines in the second step of the hyperoxidation pathway (see text).