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. 2022 Apr 5;61(8):665–677. doi: 10.1021/acs.biochem.2c00080

Pre-Steady-State Reactivity of Peptidylglycine Monooxygenase Implicates Ascorbate in Substrate Triggering of the Active Conformer

Evan F Welch †,‡, Katherine W Rush †,§, Renee J Arias †,*, Ninian J Blackburn †,*
PMCID: PMC9064607  NIHMSID: NIHMS1792198  PMID: 35380039

Abstract

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Peptidylglycine monooxygenase (PHM) is essential for the posttranslational amidation of neuroendocrine peptides. An important aspect of the PHM mechanism is the complete coupling of oxygen reduction to substrate hydroxylation, which implies no oxygen reactivity of the fully reduced enzyme in the absence of peptidyl substrates. As part of studies aimed at investigating this feature of the PHM mechanism, we explored pre-steady-state kinetics using chemical quench (CQ) and rapid freeze-quench (RFQ) studies of the fully reduced ascorbate-free PHM enzyme. First, we confirmed the absence of Cu(I)–enzyme oxidation by O2 at catalytic rates in the absence of peptidyl substrate. Next, we investigated reactivity in the presence of the substrate dansyl-YVG. Surprisingly, when ascorbate-free di-Cu(I) PHM was shot against oxygenated buffer containing the dansyl-YVG substrate, <15% of the expected product was formed. Substoichiometric reactivity was confirmed by stopped-flow and RFQ EPR spectroscopy. Product generation reached a maximum of 70% by the addition of increasing amounts of the ascorbate cosubstrate in a process that was not the result of multiple turnovers. FTIR spectroscopy of the Cu(I)–CO reaction chemistry was then used to show that increasing ascorbate concentrations correlated with a substrate-induced Cu(I)M–CO species characteristic of an altered conformation. We conclude that ascorbate and peptidyl substrate work together to induce a transition from an inactive to an active conformation and suggest that the latter may represent the “closed” conformation (Cu–Cu of ∼4 Å) recently observed for both PHM and its sister enzyme DBM by crystallography.

Introduction

Mononuclear copper monooxygenases constitute a family of metalloenzymes important to both the health- and energy-related sciences. A prime example is dopamine β-monooxygenase (DBM),1,2 which catalyzes a critical step in catecholamine biosynthesis, as does its insect homologue tyramine β-monooxygenase (TBM).3 Similarly, peptidylglycine α-amidating monooxygenase (PAM)4,5 is a bifunctional enzyme and the only known enzyme to convert glycine-extended neuropeptide hormones into their active C-terminally amidated forms. Lytic polysaccharide monooxygenases (LPMOs) contain functional mononuclear copper centers, which catalyze the chemically challenging oxidation at hexose C1 and/or C4, making them important enzymes in biomass degradation.6 Finally, new investigations into the structure and function of particulate methane monooxygenase (pMMO) denote the likelihood of a mononuclear copper site as the catalytic center for alkane hydroxylation.7,8

The catalytic core (PHMcc, residues 42–356) of PHM catalyzes the first step in peptide amidation via a copper-dependent hydroxylation of the α-C atom of the glycine-extended propeptide C-terminus (Figure 1). The reaction involves the transfer of four electrons to O2, two from the C–H bond and two from an external reductant. To achieve this conversion, the enzyme stores the two external electrons on two noncoupled copper atoms (CuH and CuM), denoted the H-site and the M-site, respectively, which are reduced to Cu(I) by ascorbate. Powerful insights into the structure and catalytic mechanism of this copper monooxygenase have been achieved in previous crystallographic,9−13 spectroscopic,14−20 kinetic,21−27 and theoretical28−30 studies. Structurally, it is known that the copper centers of WT PHMcc11,12 and its complexes with exogenous ligands9,31 are mononuclear and separated by 11 Å across a solvent-filled cleft (Figure 1). The M-site (CuM) is deemed responsible for initial oxygen binding and the ensuing catalytic chemistry10 and is coordinated by H242, H244, and a weak, yet catalytically essential interaction with the thioether of M314.14,32−34 The two reducing electrons provided to the M-site are hypothesized to be supplied by the H-site (CuH) via long-range ET.12,26,35−37 It has been reported that exogenous ligands (O2, CO, peroxide, and azide) can bind to the catalytic M-site but are excluded from the H-site by electronic or steric factors, which are incompletely understood.9,10,31,37,38

Figure 1.

Figure 1

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Top: structure of the catalytic core of the peptidylglycine monooxygenase with bound substrate di-iodotyrosyl-glycine (pdb code 3OPM). The left panel shows the protein fold architecture and the position of the two copper centers, which are separated by 11 Å. The right panel shows an expanded view of the metal centers and their coordinated residues. Copper atoms are presumed to be in the oxidized Cu(II) state. Bottom: reaction catalyzed by PHM.

A number of mechanisms have been proposed to explain how the electron is transferred across the 11 Å solvent-filled cleft that separates the H- and M-subdomains. Among these, substrate-mediated ET12 and transfer across the ordered solvent26,27 have secured the most support but another possibility proposed on the basis of crystallography2 and computational modeling30 is that the enzyme undergoes a conformational change to bring the copper sites closer together. Various crystallographic studies have shown examples of DBM2 and PHM13 in both open and closed conformations, although the biological relevance of these conformations has not yet been established. In one study, two conformations of DBM were observed in the same asymmetric unit. In one monomer, the structure of the catalytic core aligns with the open state of PHM, where the copper ions are separated by approximately 11 Å (13–14 Å in DBM), while in the second monomer, it aligns with a closed structure of PHM (mutant variant H108A in complex with citrate). In each of the closed structures, the copper sites are partially occupied (DBM) or half-filled (PHM, H-site absent), raising the question of whether the closed conformer is an artifact of copper loss. However, this closed conformation offers an appealing route for ET between the H- and M-sites and has been shown to be an energetically feasible active state by QMMM calculations.30

The likelihood of multiple conformational states of the enzyme is supported by spectroscopic studies. These studies focus on CuM binding to CO and demonstrate a substrate-induced electronic perturbation of the diatomic CO/O2 molecule.18 CO forms a complex with the ascorbate-reduced enzyme with a C≡O stretching frequency (v(CO)) of 2093 cm–1. The decrease in v(CO) from that of free CO (2143 cm–1) is consistent with back-bonding from the filled d-electron manifold into the empty π* CO antibonding orbitals and is assisted by electron-rich imidazole ligands. In the presence of the peptidyl substrate, a red shift in v(CO) is observed, with acetyl-YVG substrate dropping the frequency to 2063 cm–1. The ∼30 cm–1 decrease in v(CO) suggests a substrate-induced increase in the electron-donating power of the CuM electronic environment. Interestingly, both populations (2093 and 2063 cm–1) remain present at high acetyl-YVG concentration, suggesting two forms of the enzyme.

Here, we explore the pre-steady-state reaction of the fully reduced enzyme in the absence of a reductant and as a function of ascorbate concentration using chemical quench, (CQ) stopped-flow, rapid freeze-quench (RFQ), and Fourier transform infrared (FTIR) approaches. We show that the hydroxylation of the dansyl-YVG substrate while rapid only proceeds to <15% of the expected product based on enzyme concentration. However, this percentage increases to about 70% in the presence of ascorbate in an ascorbate-dependent manner. Pulse-chase data suggest that this is not the result of multiple turnovers but represents an activation of the Cu(I)–enzyme by excess ascorbate. Additionally, the effect appears to correlate with an increase in the species responsible for the 2063 cm–1 infrared band. Taken together, the data require a new hypothesis for substrate activation, one possibility of which includes an ascorbate-induced conformational change to bring the protein from an open to a closed conformation, facilitating electron transfer between the copper sites.

Materials and Methods

Chemicals

Buffer components and sodium ascorbate were purchased from Sigma-Aldrich with purities of >99%. The substrate acetyl-Tyr-Val-Gly (Ac-YVG) was purchased from Peptide International. The substrate dansyl-Tyr-Val-Gly (Dns-YVG) was purchased from Chi Scientific.

Cell Production and Protein Purification

PHM WT was produced from transfected CHO cells in an Accusyst Minimax bioreactor and reconstituted to 2 Cu(II) atoms per monomer. All procedures and analytical methods were as described previously.18,34,39 Protein concentration was determined using OD280(1%) = 0.980 with a Cary 50 spectrophotometer. After purification, the proteins were stored at −80 °C. Unless otherwise stated, reactions were carried out in 50 mM sodium phosphate buffer pH 7.5. Reactions were typically carried out at pH 7.5 since the single-turnover rates were not too fast to measure at this pH.

Copper Reconstitution and Analysis

The protein was reconstituted by the addition of 2.5 molar equivalents of copper sulfate by a syringe pump over the course of an hour. Excess copper was removed by dialysis in 3 L of pH 7.5 sodium phosphate buffer overnight. Reconstitution was verified using a Perkin-Elmer Optima 2000 DV inductively coupled plasma optical emission spectrometer (ICP-OES) and was determined to be 2.0 ± 0.2 Cu(II) per protein molecule (Table S1).

Generation of Ascorbate-Free Fully Reduced PHM

The fully reduced PHM was generated by the addition of 2.5 reducing equivalents (1.25 equiv per copper) of buffered sodium ascorbate as a highly concentrated small volume with mixing to a solution of anaerobic oxidized enzyme in a Vacuum Atmospheres glovebox. A color change was observed immediately from cerulean blue to pale straw. The solution was then desalted anaerobically using two passes in 7 K MWCO Zeba spin desalting columns (Thermo Fisher). The specification of these columns quotes a value of 5–10% of low-molecular-weight solute remaining after each passage, implying a negligible amount of ascorbate remaining after two passages. ICP-OES measurement of copper concentration after this protocol showed that little or no copper loss occurred during ascorbate reduction and removal. Representative data are listed in Table S1.

Oxygen Reactivity of Fully Reduced Ascorbate-Free PHM in the Absence of Peptidyl Substrate

Six hundred microliters of a 500 μM solution of oxidized PHM (1 mM in Cu(II)) was split into three portions of 200 μL each. Sample 1 was transferred directly to an EPR tube for determination of the EPR-detectable Cu(II) in fully oxidized PHM. Samples 2 and 3 were reduced anaerobically with ascorbate and made ascorbate-free as described above. Sample 2 was transferred anaerobically to an EPR tube for determination of the EPR-detectable signal in the fully reduced anaerobic ascorbate-free enzyme. Sample 3 was exposed to air with shaking for 120 s and then transferred to an EPR tube for Cu(II) quantitation. EPR spectra of all samples were measured, and the concentration of EPR-detectable Cu(II) was determined via double integration versus a 300 μM Cu(II)–EDTA standard measured under identical conditions. Spectral analysis was performed using GRAMS AI spectroscopy software (Thermo). EPR spectra were measured on a Bruker Elexsys E500 spectrometer equipped with a superX microwave bridge and a dual-mode cavity with a helium flow cryostat (ESR900, Oxford Instrument, Inc.). The following experimental conditions were used: frequency 9.63 GHz, temperature 100 K, microwave power 20 mW, gain 10 dB, modulation amplitude 10 G, and sweep time 84 s.

Fully Reduced Ascorbate-Free Single-Turnover Kinetics

The pre-steady-state reaction (single-turnover) of the fully reduced ascorbate-free PHM was interrogated using a BioLogic microvolume quench-flow instrument (QFM-4000). Six hundred microliters of fully reduced ascorbate-free PHM was preincubated with 600 μM of dansyl-YVG, preloaded into a syringe, and sealed with a septum. Sodium phosphate buffer (20 mM) was saturated with oxygen by bubbling with pure humidified oxygen gas and then loaded into a syringe sealed with a septum. The fully reduced ascorbate-free PHM + dansyl was shot against an equal volume of oxygenated buffer at varied aging times: 5, 20, 40, 60, 80, 100, 150, 200, 250, 300, and 500 ms. The reaction mixture was quenched at each aging time with an equal volume of 8% trifluoroacetic acid (TFA) and then diluted offline with 100 mM MES buffer resulting in a final concentration of 100 μM PHM + 100 μM Dns-(substrate + product). In experiments with added ascorbate, the quenched, diluted reaction mixture also contained 100 μM, 500 μM, 2 mM, or 6 mM ascorbate. Samples were analyzed via high-performance liquid chromatography (HPLC) with a Varian Pro Star solvent delivery module equipped with a Varian Pro Star model 410 autosampler (250 μL syringe, 100 μL sample loop) on a 250 mm × 4.6 mm Varian Microsorb-MV 100-5 C18 column as previously reported.35 Dansyl fluorescence was monitored with a Waters 474 scanning fluorescence detector at λex = 365 nm and λem = 558 nm. This fluorescence was used to monitor the Dns-YVG substrate concentration and hydroxylated product (Dns-YVG-OH). The product and substrate were resolved via an isocratic method using 25% solvent B (0.1% TFA in acetonitrile) and 75% solvent A (0.1% TFA in water). Substrate concentrations were determined via standard curves of known concentrations of Dns-YVG in 0.1% TFA in water.

Ascorbate Dependence of the Pre-Steady-State Reaction

The oxidized, 600 μM PHM WT was preincubated with 600 μM Dns-YVG. The PHM and Dns-YVG mixture were gently infused with pure oxygen in a 5 mL septum-fitted Wheaton vial for approximately 30 min at 23 °C. When removing the vial from the oxygen infuser, a small amount of oxygen overpressure was allowed to remain in the vial. Anaerobic ascorbate was prepared from powder in a Vacuum Atmospheres anaerobic glovebox using anaerobic buffer (50 mM NaP, pH 7.5) and was prepared in 200 μM, 1 mM, 4 mM, and 12 mM concentrations. Ascorbate mixtures were preloaded into syringes within the anaerobic chamber and sealed with septa when removing from the chamber. Ascorbate dependence was assessed by shooting the 600 μM PHM + Dns-YVG mixture against an equal volume of anaerobic ascorbate, with a set aging time of 300 ms. The samples were quenched with 8% TFA (4% final concentration) in water and analyzed via HPLC as described above.

Pre-Steady-State Kinetics at High Ascorbate Concentration

Pre-steady-state kinetics were performed using the QFM-4000 system as mentioned above but with the following adjustments: 600 μM PHM and 600 μM Dns-YVG were preincubated together and infused with pure oxygen at 23 °C. Anaerobic ascorbate was prepared in the anaerobic chamber, preloaded into a syringe, and sealed with a septum. Single-turnover kinetic curves were obtained by shooting the 600 μM PHM + Dns-YVG mixture against anaerobic ascorbate, resulting in a final concentration of 6 mM at varied aging times: 5, 20, 40, 60, 80, 100, 150, 200, 250, 300, and 500 ms. The samples were quenched with 8% TFA (4% final concentration) in water and analyzed via HPLC as described above.

“Pulse-Chase” Kinetics with Acetyl-YVG and Dansyl-YVG

To determine if the enzyme was undergoing multiple turnovers, a pulse-chase experiment was performed using the QFM. Oxidized, 600 μM PHM WT was preincubated with 600 μM Dns-YVG. The PHM and Dns-YVG mixture were gently infused with pure oxygen in a 5 mL septum-fitted Wheaton vial for approximately 30 min at 23 °C. When removing the vial from the oxygen infuser, a small amount of oxygen overpressure was allowed to remain in the vial. Anaerobic ascorbate + acetyl-YVG (Ac-YVG) was prepared from powder in a Vacuum Atmospheres anaerobic glovebox using anaerobic buffer (20 mM NaP pH 7.5) at 10 mM ascorbate plus 3 mM Ac-YVG. The ascorbate mixture was preloaded into syringes within the anaerobic chamber and sealed with septa when removed from the chamber. Repeated turnover was then assessed by shooting the 600 μM PHM + Dns-YVG mixture against an equal volume of anaerobic ascorbate Ac-YVG mixture, with aging times of 0, 300, and 1000 ms performed in triplicate. The mixture was then quenched with an equal volume of 8% trifluoroacetic acid (TFA) and then diluted with 20 mM NaP buffer resulting in a final concentration of 100 μM PHM + 100 μM Dns-(substrate + product) + 1.67 mM ascorbate + 500 μM Act-YVG. Samples were analyzed by HPLC with a Varian Pro Star solvent delivery module equipped with a Varian Pro Star model 410 autosampler (250 μL syringe, 100 μL sample loop) on a 250 mm × 4.6 mm Varian Microsorb-MV 100-5 C18 column as described above and previously reported.35

Fully Reduced Ascorbate-Free Stopped-Flow Kinetics

Ascorbate-free reduced PHM WT was prepared in the anaerobic chamber as described above. The protein was prepared in a pH 7.5 buffer system containing equal volumes of 50 mM each of MES, HEPES, and sodium phosphate. WT PHM (500 μM) was shot against 5 mM Ac-YVG + 20 mM sodium azide. Data were collected for 500 ms at 390 nm, with readings taken every 5 ms in an Applied Photosystems SX20 stopped-flow module.

Fully Reduced Ascorbate-Free Rapid Freeze-Quench EPR

Rapidly frozen EPR samples were prepared using a Quench-Flow-3 (KinTek) apparatus configured for RFQ by the manufacturer. Reduced, 200 μM ascorbate-free protein samples (400 μM in Cu(I)) in anaerobic buffer and 200 μM acetyl-YVG substrate in fully oxygenated buffer ([O2] ≈ 1.1 mM before 1:1 dilution on mixing) were loaded onto the instrument and rapidly mixed at the desired time point. A universal buffer was used comprised of equal volumes of 50 mM MES, HEPES, and CHES adjusted to pH 7.5. Mixtures were quenched in liquid ethane at −170 °C. The resulting snow was then packed into 707-SQ-250M EPR tubes (Wilmad LabGlass). Oxidation was monitored in a Bruker E500 X-Band EPR spectrometer equipped with a superX microwave bridge and a dual-mode cavity with a helium flow cryostat (ESR900, Oxford Instrument, Inc.). The experimental conditions were frequency 9.63 GHz, temperature 100 K, microwave power 20 mW, gain 10 dB, modulation amplitude 10 G, and sweep time 84 s.

Fourier Transform Infrared Spectroscopy (FTIR)

PHM (500 μM) in 20 mM pH 7.5 NaP buffer was deoxygenated and saturated with humidified CO by streaming across the surface of the solution with pure CO gas for 5 min with gentle agitation. Once carbonylated, Dns-YVG was added to a final concentration of 2.5 mM, and then a small aliquot of 2 M anaerobic buffered sodium ascorbate was added to achieve final concentrations of ascorbate between 0 and 25 mM. The mixture was then carbonylated again for 5 min. One hundred microliters of solution was then added by a syringe into the IR cell composed of two CaF2 windows separated by a 50 μm Teflon spacer. FTIR data were recorded on a Bruker Tensor 27 FTIR spectrophotometer continuously purged with CO2-free dry air as previously described.18 One thousand scans were collected for both protein sample and buffer blank from 2250 to 1900 cm–1 at a resolution of 2 cm–1. Spectral analysis including subtraction of the buffer blank was performed using GRAMS AI spectroscopy software (Thermo).

Ratios of CO bands at 2063 and 2093 cm–1 were obtained from the peak heights rather than peak integrals. The rational for this less rigorous approach stems from several factors; the cell was taken apart and reassembled between measurements taken over several days, which introduced small changes in path length and fill quality. Slight differences in protein and copper concentrations were introduced due to evaporation during carbonylation. Finally, different calcium fluoride windows were used across the experiment, which may have very slightly different optical properties.

Results

PHM Is Unreactive to Oxygen in the Absence of Substrate

Consensus mechanisms for PHM and DBM are predicated on the initial binding of oxygen to the reduced Cu(I)M-site. To probe this chemistry, we explored the sensitivity of the Cu(I)M-site to oxygen in the absence of substrate. Figure 2 shows EPR spectra of PHM as a function of time exposed to oxygen. A sample of the fully oxidized enzyme was divided into three aliquots. One aliquot was transferred to an EPR tube and measured without further modifications. A second aliquot was reduced anaerobically with 2 molar equivalents of ascorbate, and the excess ascorbate was removed by two passages through a spin desalting column and the EPR spectrum was measured. The third sample was reduced and ascorbate-depleted as described but then exposed to oxygen for 2 min, and the EPR spectrum was determined once again. The spectra were double-integrated and compared to the doubly integrated EPR spectrum of the first aliquot, which had not been reduced. The results are shown in Figure 2. Ascorbate reduction and removal resulted as expected in a low integral (3.5% EPR-detectable). However, exposure to oxygen for 2 min generated only 11.3% of the fully oxidized Cu(II) signal, indicating oxidation on a time scale much slower than the catalytic rate. While the absence of an EPR spectrum could be attributable to a spin-coupled cupric superoxo species, such an entity should be labile leading to superoxide dissociation and disproportionation, ultimately leading to an oxidized Cu(II)M-center.

Figure 2.

Figure 2

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Top: EPR spectra of WT PHM mixed with oxygenated buffer at various time points; the purple trace shows fully oxidized, the black trace shows the ascorbate-reduced anaerobic enzyme, and the red trace shows a 120 s reoxidized enzyme. Bottom: quantitation via double integration (colors are the same as for spectra) and calibration against a standard solution of 300 μM Cu(II)–EDTA (blue). EPR conditions: frequency 9.63 GHz, T = 100 K, microwave power 20 mW, gain 10 dB, modulation amplitude 10 G, and sweep time 84 s.

The low reactivity of the fully reduced ascorbate-free enzyme with oxygen is not unexpected since a notable feature of these enzymes is the tight coupling of oxygen reduction to product formation24,25,34 as well as evidence that suggests that substrate binding precedes oxygen binding.40 These data indicate that oxygen reactivity is contingent on other factors, where the reactivity and/or redox potential of the Cu(I)M-center is influenced by substrate binding. Additionally, the data demonstrate that in the absence of peptidyl substrate, exogenous ascorbate is not required to maintain the reduced state of the enzyme, even over a relatively long duration of oxygen exposure. Interestingly, while the fully oxidized enzyme clearly shows resolved components due to the two chemically inequivalent copper centers, the slow, noncatalytic reoxidation appears to favor one of these sites, as predicted by previous redox potential measurements.41

Reduction Stoichiometry Is Dependent on Exogenous Ascorbate

To further our understanding of the PHM reaction mechanism, we explored the pre-steady-state reactivity of the fully reduced ascorbate-free enzyme with oxygen and peptidyl substrate. Under these conditions, peptidyl substrate binding to the dicopper(I) enzyme should activate the catalysis and lead to rapid generation of the α-hydroxyglycyl product. The fully reduced, ascorbate-free PHM was shot against the stoichiometric dansyl-YVG substrate dissolved in buffer saturated with O2 (1.1 mM) in the absence of an exogenous ascorbate using a BioLogic QFM-4000 quench-flow microvolume mixture instrument. Here, the substrate is the limiting reagent, so the reaction should terminate when all substrates have been converted to products. Reactions were acid-quenched at successive time points with TFA, and both product and substrate were quantified via HPLC using fluorescence of the associated dansyl group. Under the canonical reaction mechanism, where ascorbate is solely the physiological electron source, this prereduced PHM should be fully primed with reducing equivalents and should be capable of reacting stoichiometrically with the substrate to reach complete product formation. Figure 3a shows the concentration of substrate and product over the course of the reaction, indicating that this reaction under ascorbate-free conditions while fast is substoichiometric, achieving only approximately 10–15% of the expected product with a kobs = 150 s–1. Despite being fully reduced, the enzyme shows only limited reactivity with substrate and oxygen. The incomplete reactivity under these conditions is also interesting given PHM’s high affinity for dansyl-YVG35 (KM = 5 μM, KD,reduced enzyme = 22 μM), suggesting that while the substrate must be binding to the enzyme in the presence of oxygen, turnover is not occurring.

Figure 3.

Figure 3

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Pre-steady-state reactivity of the fully reduced ascorbate-free PHM. (a) Time evolution of the product in the absence of excess ascorbate, kobs = 150 s–1: blue trace represents the consumption of dansyl-YVG substrate and orange trace is the rate of generation of the dansyl-α-hydroxyglycine-YVG product. (b) Time evolution of substrate consumption (blue) and product generation (orange) in the presence of excess ascorbate (6 mM), kobs = 36 s–1. (c) Bar graph showing an increase in the mole fraction of product per mole of enzyme as a function of added ascorbate. (d) Ascorbate dependence of product formation fitted to a ligand-binding event with KD = 117 μM.

We then explored the role of ascorbate as a possible reaction initiator by titrating the reactivity with increasing amounts of ascorbate. The addition of ascorbate resulted in an increase in the stoichiometry of product formation in a concentration-dependent manner (Figure 3b,c), reaching a final stoichiometry of ∼70% at an ascorbate concentration of 2 mM. Figure 3b shows that in the presence of excess exogenous ascorbate the reaction proceeds as expected with the rapid conversion of substrate to product in a manner consistent with first-order kinetics, with a pseudo-rate constant kobs = 36 s–1 that is over 4-fold slower than for the ascorbate-free enzyme. This suggests that in the presence of ascorbate some other process is rate-limiting. To address whether the inability to generate 100% product under saturating conditions of ascorbate was due to copper loss, we measured the Cu concentration after ascorbate reduction and removal (Table S1). Results show that in some samples, copper loss equivalent to 0.3 Cu/protein was detected, which could account for the 70% ceiling on product since the loss of one of the two coppers per enzyme eliminates the activity due to both coppers in that molecule.

The ascorbate dependency of the pre-steady-state reaction is shown in Figure 3c, while Figure 3d plots the mole ratio of product formation as a function of ascorbate concentration. Ascorbate is seen to influence the stoichiometry of product generation via a process that involves a binding event with a KD of 117 μM.

Ascorbate Dependency Is Not the Result of Multiple Turnovers

A plausible explanation for the observed behavior of ascorbate dependency of product stoichiometry is the hypothesis that a large population of the enzyme is unreactive, perhaps due to aggregation or some other feature of protein preparation. In this scenario, only a small population of the enzyme is functional but can undergo multiple turnovers under the conditions of the chemical quench experiment when excess reductant is supplied. To assess this possibility, a pulse-chase experiment was performed. Here, the oxidized enzyme treated with 1 equiv of dansyl-YVG in buffer saturated with O2 was shot against excess ascorbate containing a 5-fold molar excess of the unlabeled acetyl-YVG (Ac-YVG). Since the enzyme is preincubated with the fluorescent-detectable dansyl-YVG, the expectation in the case of the fully reactive enzyme is rapid pre-steady-state turnover to generate the dansyl-labeled product in near-stoichiometric amounts. In the case of a bulk-unreactive enzyme, the expectation is an approximate 5-fold reduction in dansyl-labeled product as the unlabeled substrate will compete with the labeled substrate for enzyme binding in the second and subsequent turnovers. Put another way, the acetyl-labeled substrate should act as a competitive inhibitor of dansylated product production. (This experiment works because dansyl- and acetyl- labeled substrates have similar affinity for the enzyme). Figure 4 shows that while the rate of the reaction was marginally decreased by the unlabeled acetyl peptide, its reaction progress resembles the uninhibited reaction, consistent with full enzyme reactivity. These data together demonstrate that the enzyme requires activation to achieve full reactivity by a process that is dependent in some way on ascorbate binding. This is not due to increased reductive activity since in the absence of ascorbate the enzyme is fully reduced.

Figure 4.

Figure 4

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Product generated from the reaction of dansyl-YVG with the equimolar fully reduced enzyme chased with a 5-fold excess of unlabeled Ac-YVG. Blue circles represent the product generated with equimolar dansyl-YVG and the fully reduced enzyme; purple diamonds represent the product generated when the same experiment is conducted in the presence of a 5-fold molar excess of Ac-YVG. The black dashed line represents the expected dansyl product after chasing with Ac-YVG if multiple turnovers occur.

Ascorbate Reduction Kinetics Indicate an Ascorbate Binding Site

As a final probe on the effects of exogenous ascorbate on PHM reactivity, stopped-flow was employed to determine the rate of reduction as a function of ascorbate. To accomplish this, azide was used as a reporter for the oxidized state of the enzyme. Azide is useful in these experiments as it binds only to the oxidized form of the enzyme and has a characteristic 390 nm UV–vis signal32 (Figure S1, Supporting Information). Additionally, the formation of the PHM Cu(II)–azido complex is not rate-limiting, being formed within the time of mixing (Figure S2, Supporting Information). Elimination of this signal by ascorbate therefore serves as a proxy for the rate of reduction. Figure 5 shows the rate of reduction as a function of ascorbate concentration. The reduction obeys saturation kinetics with KD = 1120 μM and kobs = 146 s–1 and indicates that ascorbate binds to the enzyme to accomplish reduction. This KD is expected to reflect the affinity of ascorbate for the oxidized enzyme during the reduction of the di-Cu(II) to di-Cu(I) states and should be similar to the apparent Km determined from the dependence of product stoichiometry on ascorbate if the latter is due merely to multiple turnovers. However, as shown in Figure 3d, the integrated peak signal for the product plotted versus ascorbate concentration yields a KD of 117 μM, an order of magnitude lower than that of the reduction event. This further supports a role for ascorbate in activation of the fully reduced enzyme over and above a simple role as a reducing agent.

Figure 5.

Figure 5

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Ascorbate dependence of the rate of PHM reduction determined by stopped-flow using the Cu(II) PHM–azido adduct (λmax = 390 nm) as a reporter for the concentration of Cu(II)–enzyme remaining at each time point. Data are fit to a pre-equilibrium binding event with KD = 1.1 mM.

Rapid Freeze-Quench EPR of the Pre-Steady-State Reaction

The above experiments demonstrate that product formation is substoichiometric without exogenous ascorbate but do not definitively identify the redox state of the enzyme during or after the pre-steady-state reaction. One possible yet unlikely explanation for the apparent ascorbate dependency is the oxidation of the enzyme without product formation, initiated by substrate and oxygen binding. Here, excess ascorbate would rereduce the enzyme to allow turnover and apparent ascorbate dependency. Such reactivity does occur in other copper monooxygenase such as the LPMOs.42 To address this possibility, rapid freeze-quench (RFQ) EPR was employed. Using this technique, the redox state of the enzyme can be assessed independent of product formation under single-turnover conditions. Figure 6 shows the results of shooting fully reduced ascorbate-free enzyme against oxygenated buffer containing 1 mole equivalent of substrate relative to the enzyme concentration. The results (Figure 6, bottom panel, and Table S2) show that in the absence of excess ascorbate negligible amounts of the enzyme become EPR-detectable after 300 ms, a time point at which product formation has ceased. This result is remarkable as consensus mechanisms predict that in the absence of reductant, turnover should generate two atoms of EPR-detectable Cu(II) for each product molecule produced. In the present experiment, if we allow for 20% product production, we anticipate 60 μM EPR-detectable Cu(II) compared with the experimentally determined amount of <3 μM. While this result should be interpreted with caution, it may suggest that an EPR-detectable entity such as a di-Cu(II) spin-coupled species is formed during or after the product is formed (vide infra). We note that a spin-coupled Cu(II)-superoxo species is less consistent with the data as any cupric superoxide formed is expected to have been converted into hydroxylated product after 300 ms.

Figure 6.

Figure 6

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Rapid freeze-quench-derived EPR spectra of the products of the pre-steady-state PHM reaction. EPR spectra were generated by rapidly mixing fully reduced ascorbate-free PHM (400 mM in Cu(I)) with the oxygenated buffer containing equimolar dansyl-YVG and freezing in liquid ethane (−120 °C). The top panel shows the actual spectra, and the bottom panel shows the quantitation of the Cu(II) signal by reference to a standard Cu(II)–EDTA sample: red, 13 ms; green, 25 ms; blue, 100 ms; orange, 300 ms; and purple, 250 μM Cu(II)–EDTA standard. To avoid errors from integration of noise peaks, intensities were determined from peak-to-peak heights at g⊥ relative to that of a 250 μM solution of Cu(II)–EDTA taken through an identical cycle of mixing, freezing, and packing. Errors in concentrations determined from the peak heights are estimated from triplicate measurements of a Cu(II)–EDTA standard to be ≅ 15% and are mainly due to differences in the packing density of the samples. EPR conditions: frequency 9.63 GHz, T = 100 K, microwave power 20 mW, gain 10 dB, modulation amplitude 10 G, and sweep time 84 s.

Cu(II)–Azido-Detected Stopped-Flow Spectrometry of the Pre-Steady-State Reaction

As discussed above, the oxidized di-Cu(II) PHM reacts with azide to generate an azide-to-copper charge-transfer complex with λmax = 390 nm. The formation of the azido complex is contingent on the oxidation of the enzyme and therefore serves as a reporter for the pre-steady state enzyme turnover from Cu(I) to Cu(II) states. Further, its rate of formation is extremely rapid, the complex being fully formed in the time of mixing in the stopped-flow (Figure S2, Supporting Information). The reaction of fully reduced, ascorbate-free enzyme with Ac-YVG and oxygen was followed using stopped-flow spectrometry monitoring the rate of appearance of the 390 nm absorption. Here, the fully reduced enzyme loaded with Ac-YVG was shot against buffer saturated with oxygen and containing 20 mM sodium azide. Figure 7 shows that in the absence of excess ascorbate less than 30% of the fully oxidized Cu(II)–azido complex is formed based on the expected absorbance for full copper oxidation (red line, Figure 7). Although this amount is larger than that based on product formation or RFQ EPR, it reinforces our findings that PHM has limited reactivity with substrate in the absence of excess ascorbate but may also suggest that product stoichiometry is influenced by other factors such as azide concentration.

Figure 7.

Figure 7

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Pre-steady-state reactivity of the fully reduced ascorbate-free PHM determined by measuring the rate of appearance of the 390 nm Cu(II)–azido adduct at 390 nm using stopped-flow spectrophotometry. 500 μM of WT PHM was shot against 5 mM Ac-YVG + 20 mM sodium azide. Data were collected for 500 ms, with readings taken every 5 ms. The top trace shows the generation of the 390 nm absorbance as a function of time in seconds. The bottom trace shows the rate curve fitted to a single exponential with kobs= 72 s–1. The red dashed line represents the absorbance expected for complete reaction where all copper is oxidized to Cu(II) determined from a separate experiment where fully oxidized Cu(II) PHM was shot against buffer containing sodium azide under identical conditions.

FTIR Indicates a Shift in Enzyme Population in Response to Ascorbate Titration

To determine the specific effect of ascorbate on PHM reactivity, we looked into insights previously garnered from FTIR. Carbon monoxide (CO) FTIR has been used extensively to probe metal oxygen-binding active sites, where it can give remarkably detailed information about structure and oxygen binding.18,37,43−45 Subtle changes to the binding site are reflected in shifts in the IR stretching frequency of the carbon monoxide. Furthermore, carbon monoxide has been shown to bind solely to the M-site in PHM and therefore it functions as a probe of M-site configuration. It has been shown previously that varied concentrations of substrates tune two separate populations of stretching frequencies.18 In the absence of substrate, the spectra are dominated by a large 2093 cm–1 stretching frequency with a minor 2063 cm–1 shoulder. Addition of substrate causes an increase in the 2063 cm–1 species with an attenuation of 2093 cm–1 in a concentration-dependent manner. Increasing substrate concentration was, however, not sufficient to eliminate the 2093 cm–1 stretch nor did a complete absence of substrate abrogate the 2063 cm–1 signal. These data suggested a pair of interconvertible states mediated by substrate binding, but the significance of these states to reactivity as well as their identity was unclear.

Peptidyl substrate’s ability to partition the enzyme population echoes ascorbate’s ability to induce a catalytically competent state, particularly since the CO binding experiments are performed in the presence of exogenous ascorbate. To probe that relationship further, we performed FTIR experiments to test if ascorbate induces an alternative enzyme configuration in a manner resembling substrate. WT PHM was exposed to carbon monoxide in the presence of substrate and varied concentrations of ascorbate. Figure 8 (top) shows that increasing the ascorbate concentration causes the ratio of 2063 to 2093 cm–1 peaks to increase in a concentration-dependent manner, reaching a maximum at 15 mM exogenous ascorbate, while Figure 8 (bottom) depicts two representative spectra at 1 and 15 mM exogenous ascorbate, respectively. Curiously, excess ascorbate beyond 15 mM resulted in a decrease in the ratio of the 2063 to 2093 cm–1 signals, but these concentrations are well above the ascorbate concentration (2 mM) required for maximum product stoichiometry. This experiment demonstrates that ascorbate, in the presence of substrate and CO, is sufficient to induce an alternate conformer of PHM.

Figure 8.

Figure 8

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Evolution of the substrate-induced 2063 cm–1 M-site Cu(I)–CO signal as a function of the added ascorbate. The top trace shows the ratio of 2063 to 2093 cm–1 peaks plotted against ascorbate concentration. The bottom trace shows the representative spectra at 1 mM (orange) and 15 mM (green).

This explicit connection between the 2063 cm–1 signal and ascorbate concentration provides insights into the role of ascorbate in catalysis. Exogenous ascorbate is here shown to both bias the population of enzyme to the 2063 cm–1 form and increase overall reactivity. This suggests that substrate and ascorbate work combinatorially to induce the 2063 cm–1 form of the enzyme and presents strong evidence that this form is the active species.

The identity of this active state remains unclear. Changes to the hydrogen-bonding network surrounding the CuM site have been proposed as one mechanism of substrate activation.13,18 Another possible interpretation is a large-scale conformational change. As discussed in more detail below, ascorbate binding could potentially induce an open-to-closed conformational transition where the Cu site separation decreases from 11 to 4–5 Å, similar to that observed in DBM. Once in proximity, electron transfer is facile between the two sites. There are several pieces of evidence that support this interpretation. First is the scale of the change: a 30 cm–1 change suggests a relatively large increase in the electron-donating power of the CuM site, larger than changes caused by mutating nearby ligands to perturb the hydrogen-bonding network. Second, the red-shifted IR frequency is reminiscent of that observed in the dinuclear arthropodal hemocyanin (2043 cm–1) and attributed to a semibridging species.44 In the context of PHM, such a bridging species is not possible unless the copper sites were in much closer proximity, as in the case of hemocyanin, which has an intracopper distance of 4 Å. Third is the observation of a closed conformer in crystal structures of the PHM H108A mutant in the presence of citrate,13 indicating that under certain conditions the interdomain closure is favored. Indeed, the ability for citrate to help induce a closed conformer is evocative, as both citrate and ascorbate are hydroxy acids. The binding of the citrate is interesting as well since it binds in a manner to bridge the CuM and CuH sites and may provide stability for the closed conformer (Figure 9).

Figure 9.

Figure 9

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Closed and open structures of DBM and PHM. Left, two conformers of DBM with Cu atom (bronze) locations modeled (PDB file 4EZL). Middle: alignment of the protein fold of the closed crystal structure of H108A in the presence of citrate (PDB file 6ALA), with the open structure of the oxidized PHM (PDB file 1PHM). PHM structures were aligned on the N-terminal (H) subdomain with an RMSD value of 0.41. The H108A-citrate and the oxidized native PHM main chains are depicted by pink and purple ribbons, respectively. To emphasize the hinge motion, the first β-strands of the M-domains are colored green (closed: H108A-citrate) and dark blue (open: oxidized PHM). Right: enlarged view of the metal binding site in the H108A-citrate PHM structure. The single copper atom of the closed H108A-citrate structure, which coordinates M-site residues H242 and H244 and H-site residue H107, is shown in pink, while the citrate molecule is shown as a green carbon backbone with oxygen atoms in red. The positions of the two copper sites in the open PHM structure are shown as transparent slate spheres.

Discussion

In the present paper, we explored pre-steady-state kinetics of PHM as a function of ascorbate concentration using chemical quench and rapid freeze-quench studies of the fully reduced ascorbate-free enzyme. When ascorbate-free di-Cu(I) PHM was shot against oxygenated buffer containing the dansyl-YVG substrate and the dansylated product quantified by HPLC using fluorescence detection, <15% of the expected product was formed. This result was recapitulated to varying degrees by RFQ EPR spectroscopy and stopped-flow monitoring of the oxidized Cu(II) species using the 390 nm Cu(II)–azido absorption as readout. Product generation could be increased to a maximum of 70% by the addition of increasing amounts of the ascorbate cosubstrate. This phenomenon could be the result of multiple turnovers of a small fraction of the “active” enzyme, with the remainder of the sample in a nonfunctional state (case 1) or activation of the fully reduced enzyme by excess ascorbate (case 2). To test which of these scenarios was most probable, we performed a pulse-chase experiment, wherein the enzyme was incubated with 1 equiv of dansyl-YVG in oxygenated buffer and then shot against a 5-fold excess of Ac-YVG. Case 1 is predicted to generate no more than 20% of the fluorescently labeled product since after the first turnover the unlabeled Ac-YVG would efficiently outcompete the labeled substrate, whereas case 2 is predicted to generate higher amounts of the labeled product. The latter scenario was observed with 50–60% of the dansylated product formed. This allowed us to exclude multiple turnovers as an interpretation of the data and to conclude that the cosubstrate ascorbate not only reduces the Cu(II) centers but also induces a conformational change to an active state. Further, the observed rate in the presence of ascorbate is 4-fold slower than that of the ascorbate-free conformer, indicating that ascorbate induces some process such as a conformational change, which becomes rate-limiting. In previous work, we had documented that the peptidyl substrate induces a downshift in the ν(CO) of the carbon monoxide complex at the M-center from 2093 to 2063 cm–1 and had suggested that this species was a substrate-triggered active conformer. Here, we found that this conformer was also induced by increasing concentrations of ascorbate, confirming that ascorbate does more than act as a reducing agent. The totality of the data led to the hypothesis that ascorbate and peptidyl substrate work together to induce a transition from the inactive to an active conformation.

The present study demonstrates a correlation between the intensity of the 2063 cm–1 CuM–CO band and the amount of product formed during pre-steady-state turnover (since both increase with increasing ascorbate), and one may cautiously infer that the species responsible for the 2063 cm–1 band is derived from a catalytically active form, where the CO ligand is bound in the same configuration as the reactive oxygen molecule. Assigning the chemical nature of the CO-bound species is therefore important for understanding the catalytic mechanism and the mechanism of substrate activation. The 30 cm–1 red shift of the CO frequency implies a considerable degree of additional back-bonding into ligand π* orbitals, which, in turn, weakens the CO bond via decreasing the bond order, and similar electronic perturbation applied to a Cu(I)–O2 complex would result in significant activation of the metal–dioxygen complex.

Downshifted frequencies in Cu(I)–CO complexes can potentially arise from a number of sources such as (i) increase in the N-donor (imidazole) coordination number, (ii) strong H-bonding between the distal NH of the imidazole of a His ligand and a protein-derived negative charge or dipole, (iii) interaction of the distal O of CO with a positive charge or dipole, and (iv) a bridging or semibridging structure where the CO can interact with a second metal ion. In model compounds, the substitution of heterocyclic rings (imidazole, pyrazole) by electron-releasing groups leads to decreased frequencies, while substitution by electron-withdrawing groups leads to increased frequencies. Table S3 of the Supporting Information lists examples that validate these trends. In a previous study,18 we showed that the substitution of Met314 by His resulted in an 18 cm–1 downshift (2075 cm–1) in the absence of substrate, but the addition of substrate led to a further downshift of 24 cm–1 (2051 cm–1). Therefore, simple substitution of the weakly binding Met by His does not account for the observed effect. Similarly, changes in the H-bonding networks, which anchor the M-site coordinating His ligands, are unlikely to generate sufficient perturbation. For example, in recent unpublished work, we perturbed the H242–Q272 H bond via a Q272E mutation and detected a mere 4 cm–1 downshift. A substrate-induced interaction of the CO distal O with a positive dipole as documented by Spiro and co-workers46 for heme carbonyls could lead to significant downshifted frequencies, yet it is hard to see how binding of either Ac-YVG or ascorbate could increase the positive charge in the vicinity of the bound CO since both carry a negative charge at the pH of our experiments. Indeed, the salt-bridge formed between the peptide carboxylate and R240 would remove a source of positive charge (protonated guanidinium) that could potentially interact with a coordinated CO. We are therefore left with the final possibility that a conformational change leads to the formation of a semibridging CO complex where the CO can interact with a second metal ion. It is worth noting that for the M314H Cu(I)–CO the measured ν(CO) of 2051 cm–1 is one of the lowest frequencies ever observed for a Cu(His)3 ligand set (Table S3) and is in the range observed for the CO complexes of hemocyanins 2063–2043 cm–1, which have the same His3 coordination44 and where the interaction of the O atom of the CO ligand with the positive charge of the second Cu(I) was deemed the most likely origin of the lowered frequency.

Red-shifted IR frequencies are also observed in the CuB–CO complexes of cytochrome-c-oxidase47,48 between 2060 and 2036 cm–1. Like the hemocyanin case, Cu(I)–CO frequencies at or below 2050 cm–1 appear to correlate with the presence of a second positively charged metal ion in the vicinity of the CO binding site. In the case of cytochrome oxidase, the Cu(I)–CO species are formed by the photodissociation of the CO ligand from the ferrous heme-a3 with subsequent rebinding to copper. Recent X-ray crystallographic studies49 have captured an X-ray photodissociated state in the crystal where the CO binds side-on to Cu(I) with Cu–C and Cu–O distances of 2.5 and 2.3 Å, respectively. While it is not known whether this state is representative of the photodissociated CuB–CO complexes detected by IR, it is indicative of a plurality of CO binding motifs that are accessible in dinuclear metal environments. In the substrate-induced PHM CuM–CO complex, we may cautiously infer that the interaction of the O atom with the positive charge of a second copper atom is an attractive interpretation of the red shift, which would imply that activation is the result of an open-to-closed conformational transition.

Our data provide experimental context for recent computational work, which has documented the energetic feasibility of the open-to-closed conformational transition in catalysis. The new theoretical computations that consider the closed conformer as the active species have suggested a binuclear intermediate, which significantly lowers the calculated activation energy for HAA from the peptide substrate.30 The new modeling shows that hydrogen atom abstraction by a cupric superoxo from the reductant ascorbate is always more favorable than from the peptide Cα–H, implying that in the presence of excess ascorbate, any cupric superoxo that forms will always rapidly form a cupric hydroperoxo and the ascorbyl radical. The study suggests that the Cu(II)M hydroperoxo species (formed only in the presence of ascorbate) drives the conversion of open-to-closed conformer which are separated by only 2 kcal mol–1, and forms a μ-oxo-, μ-hydroxo-mixed-valence Cu(I)–Cu(II) entity as the active intermediate. While experimental data are yet to support such an intermediate, there is clearly sufficient evidence to warrant re-examination of the canonical mechanism. The extremely small amount of EPR-detectable Cu(II) observed in our RFQ pre-steady-state experiment offers the intriguing possibility that the closed conformer may persist after product formation as a starting point for subsequent catalytic cycles.

An open-to-closed mechanism is also more consistent with a number of experimental observations that do not fit well with the consensus mechanism. The latter posits that the initial attack of O2 on the reduced CuM center generates an electrophilic Cu(II)–superoxo intermediate capable of H atom abstraction from the peptidylglycine Cα to form a substrate radical. This mechanism fails to explain (i) how substrate binding converts the redox-inactive Cu(I)M-site to an active state apparently associated with the 2063 cm–1 Cu(I)M–CO species; (ii) the requirement for methionine at position 314 and the inactivity of the M314H variant18 since Cu(I)His3 systems are known to be reactive toward O2 in inorganic model compounds;50 and (iii) how substrate hydroxylation can be driven by hydrogen peroxide from the oxidized Cu(II)–enzyme (peroxide shunt) in a slow catalytic reaction in which the 18O of labeled peroxide is scrambled 60–70% with ambient 16O2 in air.35 However, an open-to-closed mechanism rationalizes all of these observations and can be discussed in the context of the H108A-citrate structure shown in Figure 9. We suggest that the peptide and ascorbate together induce the open-to-closed transition in a rate-limiting step to form the active “binuclear” state. This is reactive to both oxygen in the Cu(I) state and hydrogen peroxide in the Cu(II) state. An interesting feature of the H108-citrate structure is the lengthening of the CuM to M314 Cu–S bond from 2.3 to 5.6 Å. This movement requires a labile fluxional Cu–S interaction that is provided by the Cu(I)–thioether bond as recently demonstrated in our studies of the SMet and SeMet complexes of a PHM CusF model32,33 but conversely would be frustrated in the M314H variant due to the stronger sigma donor interaction of the Cu(I)–N(H314) bond.

The peroxide shunt chemistry is also easily interpreted within the binuclear formalism. From work published nearly two decades ago,35 we showed that the reaction was catalytic (at least 40 turnovers) and was 100% coupled to peroxide consumption. The conclusion that a mononuclear Cu(II)–peroxo species must be a reactive intermediate was negated by the observation that 60% of the 18O label was scrambled with ambient 16O2 in air when H218O2 was used as the source of peroxide, implying the formation of an intermediate that was in equilibrium with ambient dioxygen at some stage of the catalytic mechanism. While difficult to rationalize within the context of the “open” PHM conformation, a binuclear intermediate formed within the closed conformer can readily explain the data due to the well-known equilibrium between di-Cu(II)–peroxo and di-Cu(I)–O2 species.

Our studies show that ascorbate appears to play a dual role in PHM catalysis both as a reductant and as an activator but the stoichiometry and binding site(s) remain obscure. While ascorbate is technically able to supply two electrons, earlier work on DBM established that it functioned as a 1-electron reductant generating the semidehydroascorbate radical as the primary product, which subsequently disproportionates to ascorbate and dehydroascorbate.51 Thus, we anticipate that during the reductive cycle, ascorbate must bind and its oxidized radical must dissociate twice to generate the fully reduced dicopper(I) enzyme. This process appears to have a KD of around 1.1 mM. On the other hand, the activation of catalysis occurs with a lower KD (117 μM), suggesting a different mode/site of ascorbate interaction with the enzyme. Yet, another study reported inactivation by ascorbate with a KD of 5 mM,52 whereas crystallography failed to identify any ascorbate binding site.4 Interestingly, the ascorbate dependence of the 2063 cm–1 CO conformer shows a maximum between 10 and 15 mM ascorbate suggestive of both activation and subsequent inactivation as ascorbate concentrations increase. The data suggest complex activation/inactivation behavior for the ascorbate cosubstrate over a concentration range of 0.1–10 mM, and it is possible that this may relate to the physiological regulation of hormone production in the vesicle where the ascorbate levels also span this concentration range.53

Notwithstanding the growing evidence for the open-to-closed mechanism, an alternative mechanism for substrate triggering in the open-only formalism may be suggested based on the calculations of Cowley and Solomon, who concluded that the formation of the Cu(II)–superoxide at the M-site was energetically favorable only if the dioxygen displaced a hydroxide ligand from the Cu(I) center.29 While Cu(I)–OH complexes are rare, crystal structures of the reduced enzyme do show water/hydroxide density near the CuM center, which appears to be a coordinated solvent species. It is therefore possible that substrate and/or ascorbate binding to the Cu(I) state deprotonates the coordinated water to a hydroxide, which destabilizes the Cu(I)M-site and induces reaction with oxygen. However, the fact that the catalytic rate is inversely proportional to pH in the 4–8 pH region is inconsistent with M-site water deprotonation as the driver of substrate triggering. We continue to search for conditions where closed conformers can be formed and studied as potential intermediates.

Acknowledgments

This work was supported by a grant from the National Institutes of General Medical Sciences (R35 GM136239) to N.J.B.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.biochem.2c00080.

  • Two figures describing an oxidized PHM titration with azide and its rate of formation studied by stopped-flow; and three tables describing the quantitation of EPR data and CO stretching frequencies for a selection of protein and model Cu(I)–CO complexes (PDF)

Accession Codes

Peptidylglycine monooxygenase copper-dependent oxidoreductase (Rattus norvegus) UniProtKB-P14925 (AMD_RAT).

Author Present Address

∥ Materials & Structural Analysis Division, Thermo Fisher Scientific, Hillsboro, Oregon 97124, United States

Author Contributions

E.F.W. performed the majority of data collection and analysis; R.J.A. assisted in cell culture, protein isolation, and in the stopped-flow measurements as well as providing mentorship to E.F.W. K.W.R. performed the RFQ data collection and assisted with EPR analysis. N.J.B. and R.J.A. designed and supervised the experimental program, and N.J.B. secured the funding.

The authors declare no competing financial interest.

Supplementary Material

bi2c00080_si_001.pdf (132.8KB, pdf)

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