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REDOX Cycling, pH Dependence, and Ligand Effects of Mn(III) in Oxalate Decarboxylase from Bacillus subtilis Umar Tariq Twahir, Andrew Ozarowski, and Alexander Angerhofer Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.6b00891 • Publication Date (Web): 31 Oct 2016 Downloaded from http://pubs.acs.org on November 4, 2016

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REDOX Cycling, pH Dependence, and Ligand Effects of Mn(III) in Oxalate Decarboxylase from Bacillus subtilis Umar T. Twahir,a† Andrew Ozarowski,b and Alexander Angerhofer a* a

Department of Chemistry, University of Florida, Gainesville, Florida 32611-7200, USA.

b

National High Magnetic Field Laboratory, 1800 E. Paul Dirac Dr., Tallahassee, FL 32310, USA

Funding Source Statement: Funding for this work was provided by the National Science Foundation under Grant CHE-1213440. Part of this work was conducted at the NHMFL, which is funded by the NSF through a Cooperative Agreement DMR 1157490, the State of Florida and the US Department of Energy. Corresponding Author: Alexander Angerhofer Fax: +1 3523920872 Phone: +13523929489 E-mail address: [email protected] Physical Address: 318A, Chemistry Lab Building P.O.Box. 117200 Gainesville, FL, 32611

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Keywords: electron paramagnetic resonance, parallel mode, oxalate decarboxylase, redox, Mn(III).

Abbreviations: B. subtilis: Bacillus subtilis; EPR: electron paramagnetic resonance; IMAC: immobilized metal affinity chromatography; OxDC: oxalate decarboxylase; OxOx: oxalate oxidase; WT: wild type


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ABSTRACT This contribution describes EPR experiments on Mn(III) in oxalate decarboxylase of Bacillus subtilis, an interesting enzyme that catalyzes the redox-neutral dissociation of oxalate into formate and carbon dioxide. Chemical redox cycling provides strong evidence that both Mn centers can be oxidized although the N-terminal Mn(II) appears to have the lower reduction potential and is most likely the carrier of the +3 oxidation state under moderate oxidative conditions in agreement with the general view that it represents the active site. Significantly, Mn(III) was observed in untreated OxDC in succinate and acetate buffers while it could not be directly observed in citrate buffer. Quantitative analysis showed that up to 16% of the EPR visible Mn is in the +3 oxidation state at low pH in the presence of succinate buffer. The fine structure and hyperfine structure parameters of Mn(III) are affected by small carboxylate ligands that can enter the active site and have been recorded for formate, acetate, and succinate. The results from a previous report by Zhu et al. (2016) Biochemistry 55, 429-434 could therefore be reinterpreted as evidence for formate-bound Mn(III) after the enzyme is allowed to turn over oxalate. The pH dependence of the Mn(III) EPR signal compares very well with that of enzymatic activity providing strong evidence that the catalytic reaction of oxalate decarboxylase is driven by Mn(III) which is generated in the presence of dioxygen.


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Oxalate Decarboxylase (OxDC) is one of three known oxalate-catabolizing enzymes and has been found in bacteria and fungi.1-4 To date the best studied OxDC is that from Bacillus subtilis, a soil bacterium which is thought to play a critical role in the biogeochemical carbon cycle.5 OxDC appears to be part of an enzymatic cascade in this prokaryote to control cytoplasmic pH when the organism encounters low pH in soil or rotting vegetation.4 B. subtilis OxDC is a bicupin enzyme that coordinates a mononuclear Mn(II) in each of its domains.6-8 It catalyzes the redox-neutral dissociation of oxalate producing one equivalent each of carbon dioxide and formate while raising the pH using molecular dioxygen as a co-catalyst.9, 10 Interestingly, the enzyme functions as an oxidase in approximately 0.2% of turnovers, thereby producing two equivalents of carbon dioxide and hydrogen peroxide.4, 9, 10 OxDC displays a strong pH dependence of its catalytic efficiency between pH4 and pH6, with the highest activity at or below pH4 and becoming inactive at pH values above 6.11, 12 Scheme 1 depicts the mechanism that has served as the main hypothesis for OxDC catalysis for many years.12-14


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Scheme 1: Currently accepted hypothesis for the catalytic mechanism by which OxDC converts the oxalate monoanion to carbon dioxide and formate.12-14 The scheme starts with a superoxide species bound to Mn(III) in the active site, presumably the N-terminal Mn. At low pH where the enzyme exhibits maximum efficiency, superoxide is most likely protonated which raises the potential of the O2/ O2·− couple and may make dioxygen a more favorable electron sink able to withdraw an electron from protein-bound Mn(II).15 At present there is no direct experimental evidence for oxygen binding to the Nterminal Mn even though it is presumed to be the catalytically active site. Nitric oxide, a structural analog of dioxygen, has been shown to be a reversible inhibitor of OxDC but does not associate with the N-terminal Mn(II) suggesting that neither of them bind there.16 However, dioxygen is needed for turnover10 and it is generally assumed to prime the Mn ion in the Mn(III) oxidized state responsible for catalysis. Based on the pH dependence of catalytic activity the substrate is presumed to be the oxalate monoanion.12 It would likely bind mono-dentate if


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oxygen binds to the same metal ion as suggested by a report showing mono-dentate binding of oxalate to a Co-substituted ∆E162 mutant of OxDC.17 Recently, Zhu et al. suggested that oxygen binds after substrate coordination to Mn(II) in wild-type OxDC based on the observation of Mn(III) by parallel mode X-band EPR only after turnover has started.18 Based on kinetic isotope effect measurements it was proposed that the rate limiting step is an initial proton coupled electron transfer step taking an electron from the substrate and generating a Mn(II) bound superoxide.7, 12, 14, 19 Superoxide production during turnover was indeed observed by spin trapping using both EPR and mass spectrometry.20 The initial electron transfer step produces an oxalate radical intermediate which is susceptible to rapid decarboxylation21 and loss of CO2 through a migration pathway leading from the N-terminal active site to the outside of the protein.22, 23 At this point a carbon dioxide radical anion intermediate, CO2·−, remains in the active site and the decarboxylase and oxidase pathways fork. Oxidase activity happens if a second electron is removed from this intermediate through an inner or outer shell mechanism resulting in hydrogen peroxide production while decarboxylase activity requires a second proton coupled electron transfer where the intermediate gets reduced and a proton placed on the carbon atom.4, 24 Zhu et al. suggested that oxidase activity is based on solvent access to the active site and it may thus be an outer-shell mechanism.17, 18 This idea is in agreement with earlier EPR spin trapping experiments on the CO2·− radical that explained oxidase activity by loss of the intermediate from the enzyme during turnover followed by rapid reaction with dissolved dioxygen.20, 23 Enzymatic activity is clearly dependent on the presence of dioxygen which is the ultimate electron sink available for the initial 1-electron oxidation of the substrate. Complete removal of dioxygen appears to be difficult suggesting that a fraction of the enzyme contains strongly bound


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dioxygen based upon 16-20% residual activity after treatment with glucose/glucose oxidase.10 Treatment with dithionite shuts down activity completely but this experiment was considered inconclusive since re-equilibration of the sample with atmospheric oxygen restored only 18% of the activity suggesting an irreversible effect of dithionite on the enzyme.10 Oxygen is presumably bound to one of the Mn-centers in OxDC and would be expected to form a Mn(III)-superoxo complex (as seen in scheme 1) which could provide the driving force for the initial oxidation step. It was previously shown that the catalytically active species in barley oxalate oxidase (OxOx) is Mn(III) which is present in the resting state of the enzyme at a fraction of only 17 to 26% of the available Mn.25 A similar suggestion regarding OxDC was made by Bornemann based on quantitative EPR of the Mn(II) recovered after chemical reduction (approximately 20%)10 and the quantitative analysis of the low-temperature CD spectrum.13 Both observations suggested that approximately 15 – 20% of the Mn in WT OxDC is in the +3 oxidation state under normal conditions.13 Direct evidence for the presence of Mn(III) by EPR was first provided by Chang et al. using parallel mode EPR with a sample that was flash-frozen under turnover conditions at a relatively high pH of 5.2 and then stored at 77 K for an extended time.26 More recently, Zhu et al. showed parallel mode X-band EPR evidence of Mn(III) at low pH under turnover conditions using citrate buffered enzyme.18 These workers suggested that oxalate binding precedes oxygen binding which is necessary to lead to observable Mn(III). Curiously, a rapid freeze quench experiment that was reported in the supplementary information of the same paper did not show any detectable Mn(III) upon mixing which raises the question whether the Mn(III) that was observed is a true intermediate or accumulates over time in a side reaction or due to increasing product concentration.


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The major difficulty in studying integer spin species, such as Mn(III) having an S = 2 total spin (non-Kramers system) is the very large zero-field splitting parameter |D| which typically exceeds the quantum energy of X-band photons. Using a parallel mode cavity allows one to apply the polarization of the microwave B1 field parallel to the external magnetic B0 field facilitating the observation of integer spin species such as high-spin Mn(III).27, 28 Parallel polarization EPR has a ∆mS = 0 selection rule which allows the observation of transitions between relatively closely spaced ms = ± 2 sublevels under low field conditions due to mixing of the Zeeman terms by the zero-field Hamiltonian.29-31 In this contribution we provide the EPR spectroscopic signature of Mn(III) after chemical oxidation of wild-type OxDC using parallel mode EPR in the X-band in the presence of small carboxylate anions. Contrary to Zhu et al.’s report18 we also detect the presence of Mn(III) in the WT enzyme at low pH in the absence of substrate when succinate or acetate are used as buffers. A quantitative analysis suggests that up to 16% of the EPR visible Mn is in the +3 oxidation state under those conditions which is in good agreement with Bornemann’s earlier results.10, 13 An explanation of this interesting buffer effect is given based on the ability of the smaller carboxylate anions used as buffer molecules (succinate and acetate) to coordinate to the Nterminal Mn. The coordination of an additional carboxylate group may affect the Mn(II)/Mn(III) potential and stabilize the +3 oxidation state on the active site Mn ion. We also show that the Mn(III) observed by Zhu et al. in OxDC in the presence of oxalate under turnover conditions is due to accumulated product, formate. High-field EPR of redox cycled preparations suggest that the N-terminal Mn(II) gets oxidized more readily than the C-terminal one which indicates that Mn(III) is located on the N-terminal site under low pH turnover conditions. The high-field experiments on oxidized high pH OxDC also revealed a carbon-based radical with g-tensor


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parameters matching well reported literature values for tyrosyl radicals. We also measured the pH dependence of the Mn(III) EPR signal using a new enzyme immobilization protocol that allowed us to use the same sample for several pH steps.32 These experiments suggest that it is the Mn(III) concentration that determines the pH dependence of catalysis and not the second pKa of oxalic acid.


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Materials and Methods Materials. The following chemicals were purchased from Fisher Scientific (Pittsburg PA) (ACS Grade) and used as received without further purification: sodium citrate, sodium formate, sodium chloride, sodium phosphate, sodium acetate, sodium hydroxide, succinic acid and tris base (2-amino-2-hydroxymethyl-propane-1,3-diol). Potassium hexachloroiridate, sodium ascorbate, bis-tris (2-bis(2-hydroxyethyl)amino-2-(hydroxymethyl)-1,3-propanediol), and piperazine were purchased from Sigma Aldrich (St. Louis, MO). All solutions were prepared utilizing 18 MΩ distilled water generated by a Thermo Scientific Barnstead Nanopure Model 7134. Protein Expression. Expression and purification of recombinant His6-tagged B. subtilis wild-type OxDC was carried out following previously published procedures.7, 8, 33, 34 To remove dissolved metals from the preparation, Chelex 100 resin (Bio-Rad, Hercules CA) was added to the enzyme after the serial dialysis steps. The solution was shaken on ice for approximately 1 h following removal of the resin. The enzyme solution was then concentrated using Amicon Centriprep YM-30 centrifugal filter units (EMD Millipore, Billerica, MA). Concentrated enzyme samples (approximately 40 mg/mL) were stored as 200 µL aliquots in Eppendorf tubes at -80oC until used for experiments. Protein concentration was measured utilizing a Bradford assay.35 This commonly used assay is based on the color change of Coomassie Blue dye upon interaction with protein. Coomassie Blue dye (Thermo Scientific Bradford Assay Kit) is normally a brown-red color, but it turns blue when it binds to protein through apparently non-specific interactions. The absorbance of the solution at 595 nm can be related linearly to the concentration of protein in a


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given sample, using bovine serum albumin (BSA, Thermo Scientific) to create a standard curve. ICP-MS determinations of metal content were performed at the University of Georgia Center for Applied Isotope Studies Chemical Analysis Laboratory (Athens, GA), and were found to typically contain 1.6 Mn/monomer in our preparations. Enzymatic Assay. The Michaelis-Menten parameters of the decarboxylase activity were determined through a coupled end-point assay measuring the production of formate, as described in previous publications.8, 10, 33 WT OxDC is allowed to react in the presence of oxalate at the desired pH for 60 seconds at 25°C, after which point the reaction is quickly quenched with 1 M NaOH. The products of this reaction are then mixed with formate dehydrogenase and NAD+, leading to the conversion of formate to carbon dioxide and the reduction of NAD+ to NADH. The absorbance of NADH at 340 nm was then measured and related to the final formate concentrations using a standard curve derived from known formate concentrations. Reaction rates were calculated by dividing the amount of formate produced by the 60-second reaction time, and accounting for dilution effects. Kinetic parameters for the WT enzyme were found to be: Km = 12 ± 3 mM, and kcat/Km = 13000 ± 3000 M-1s-1, matching well with previous literature reports.7, 8 EPR Spectroscopy. EPR experiments were performed on a Bruker ELEXSYS E580 CW/Pulsed or a Bruker ELEXSYS-II E500 CW X-band spectrometer equipped with a dual mode cavity (Bruker ER-4116DM). Experimental conditions were typically: 100 kHz modulation frequency, 10 G modulation amplitude, 0.63 mW microwave power, and sample temperature set to 5 K. High field/frequency measurements were collected on a home-built variable frequency/field broadband transmission homodyne spectrometer at 406.4 GHz in a field ranging from 13.9 T to 14.5 T, 50 kHz modulation frequency, 1 G modulation amplitude, 0.2 mT/s sweep


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rate, and in a temperature range of 3 to 20 K.36 Simulations of experimental spectra were carried out using the EasySpin toolbox for MATLAB.37 Enzyme Oxidation. After expression and purification of the enzyme it was exchanged into a buffer consisting of 50 mM succinate, acetate, or citrate, 500 mM NaCl, for low pH experiments, or 50 mM potassium phosphate, 500 mM NaCl, for experiments at pH 8.0. Prior to exchanging the protein into the appropriate carboxylic acid buffers, it was necessary to lower the pH which was carried out in a poly-buffer containing 50 mM tris, 50 mM bis-tris, 50 mM piperazine, 50 mM succinate, 500 mM NaCl, and 20% glycerol. The protein was dialyzed into the buffer overnight and then pH adjusted in 1 pH unit decrements to the desired lower pH. Once the final pH was reached, the sample was then dialyzed into its final buffer. Stocks of 0.1 M hexachloroiridate and ascorbate in the respective buffer and pH matching that of the protein from which 2 to 10 µL aliquots were added to 200 µL of enzyme solution for stepwise oxidation and reduction experiments.


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Results Redox Cycling of OxDC. Chemically oxidized enzyme was prepared by step-wise addition of the mild oxidant hexachloroiridate(IV) up to a concentration of 8 mM at low pH (4.2) and up to 6 mM at high pH (8.0). These amounts were needed to convert OxDC to a saturated oxidation state where additional increase of the amount of oxidant did not visibly change the EPR spectrum anymore. Figs. 1 and 2 show the EPR results for this experiment in both parallel and perpendicular modes at X-band.

Figure 1: X-band EPR spectra of WT OxDC at pH4.2. Left panel (A): perpendicular mode, right panel (B): parallel mode. Blue trace: Enzyme in succinate buffer as prepared, red trace: after cumulative addition of 8 mM hexachloroiridate(IV), green trace: after addition of 8 mM ascorbate. All spectra were taken on the same sample at 5 K. Subsequent additions of oxidant and reductant were made after the sample was thawed. EPR parameters: frequencies for perpendicular and parallel mode were 9.618 GHz and 9.331 GHz, respectively, modulation frequency was 100 kHz, modulation amplitude was 10 G, and microwave power 0.63 mW.


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Figure 2: X-band EPR spectra of WT OxDC at pH8.0. Left panel (A): perpendicular mode, right panel (B): parallel mode. Blue trace: Enzyme in phosphate buffer as prepared, red trace: after cumulative addition of 6 mM hexachloroiridate(IV), green trace: after addition of 6 mM ascorbate. All spectra were taken on the same sample at 5 K. Subsequent additions of oxidant and reductant were made after the sample was thawed. EPR parameters: frequencies for perpendicular and parallel mode were 9.618 GHz and 9.331 GHz, respectively, modulation frequency was 100 kHz, modulation amplitude was 10 G, and microwave power 0.63 mW.

In each case the experiment started with the enzyme as prepared (see Materials and Methods) either at low pH4.2 in 50 mM succinate buffer, or at high pH8.0 in 50 mM phosphate buffer. Initially we increased the oxidant concentration in increments of 1 mM and monitored the disappearance of the Mn(II) signal and the appearance of the oxidized state (see supporting information, figs. S1 – S4). This required repeated freezing and thawing of the same sample. Addition of oxidant was performed with small 2 µL aliquots of concentrated stock in order to keep the dilution effect at a minimum. Since there was no noticeable change in the EPR spectra after an oxidant concentration of 8 mM Ir(IV) at low pH and 6 mM at high pH we assumed that the majority of the Mn(II) ions were oxidized under these conditions. The experiments in figs. 1


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and 2 were performed using a Bruker dual mode cavity that permits to perform both perpendicular and parallel mode EPR on the same sample by selecting two different cavity modes. At the end of the experiments on oxidized samples, they were thawed on ice and then reduced with a small aliquot of concentrated ascorbate solution, refrozen, and their EPR spectra taken again.

Comparison of the perpendicular mode EPR spectra shows a drastic decrease in the Mn(II) signal upon oxidation (compare red with blue trace) at both pH values. At low pH the Mn(II) signal is diminished to less than 10% of its original intensity while at high pH it has completely vanished. The spectrum of the oxidized sample at high pH shows a strong carbonbased radical signal at geff ≈ 2 (truncated along the y-axis in fig. 2A). It appears similar to the tyrosyl radical that was observed to accumulate under turnover conditions in earlier experiments (vide infra).26 No such radical is observed in the low pH experiment.

Focusing now on the parallel mode spectra, we note that a Mn(II) signal in virgin WT OxDC appears in the spectral range between 1000 and 1800 G centered near geff ≈ 5.2. It is most likely due to a ∆mS = ±2 transition of Mn(II) and shows the same hyperfine splitting with the 55

Mn nucleus as the perpendicular mode spectrum near geff ≈ 2, i.e., A = 249 MHz (88.5 G). Its

appearance is a convenient way to monitor the presence of and quantitate Mn(II) in the sample when using parallel mode. After oxidation with iridate the Mn(II) trace almost completely vanishes. The low pH sample still contains a few broad lines in the same field range while the high pH spectrum shows a more or less flat line between two positive peaks of a residual spectrum that can be traced back to the presence of paramagnetic oxygen in the sample (see supporting information, fig. S5). Upon reduction of the low-pH sample with ascorbate the Mn(II)


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spectrum reappears. Its intensity is slightly lower than in the original sample which is due to the dilution effect by the aliquots of oxidant and reductant that had been added to the sample.

Interestingly, in the high-pH experiment the final Mn(II) EPR intensity at geff ≈ 2 (perpendicular mode) and geff ≈ 5.2 (parallel mode) is higher than in the starting sample even though it should be the same if ascorbate completely reverses the effects of hexachloroiridate treatment. This may be due to a shift in EPR intensity from a very broad spectrum (C-terminal site with large |D|) to a narrow spectrum (N-terminal site with small |D|) during the course of the experiment, essentially superposing the spectra of both sites in the geff ≈ 2 and geff ≈ 5.2 regions after reduction with ascorbate. It may indicate a conformational change in the high-pH protein as it undergoes the oxidation-reduction cycle changing the C-terminal Mn(II) from penta- to hexacoordinate.

Magnetic Parameters of Mn. Turning our attention back to figure 1B, we notice a very strong sextet signal in the field range between 600 and 1000 G, centered near 760 G (geff ≈ 8.8) for the low-pH oxidized sample. This is the field range where parallel mode EPR for the nonKramers high-spin Mn(III) ion in a reasonably rhombic environment is to be expected.27, 39-45 The high-pH sample in fig. 2B does not show this sextet signal after oxidation but instead exhibits a broad trough in the EPR trace, centered around 700 G (geff ≈ 9.5). This signal may perhaps be due to a different ligand environment for Mn(III) with a distorted parallel-mode spectrum broadening out the distinct 55Mn hyperfine splitting. Since the enzyme is not active at high pH we will not focus on this high pH signal further. Fig. 3 shows simulations of the parallel mode Mn signals for the low-pH form of WT OxDC. The geff ≈ 8.8 signal of Mn(III) is shown in


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fig. 3A after the sample had been oxidized. The geff ≈ 5.2 signal in fig. 3B is from the untreated WT OxDC sample. Simulations were carried out with the Easyspin toolbox for Matlab™.37

Figure 3: Parallel-mode X-band EPR spectra and spectral simulations of Mn(III) (A, left panel), and Mn(II) (B, right panel). Experimental spectra are in blue, the simulations in orange.

The magnetic parameters obtained from the simulation of the Mn(II) signal are: g = 2.001, A = 249 MHz, and |D| = 0.0447 cm-1, |E/D| = 0.17. These values agree with the expected magnetic parameters for the low pH N-terminal Mn(II) form reported by Tabares et al.38 The Mn(III) spectrum in fig. 3A could be well represented by a simulation using g = 2.000, A = 140 MHz, D = -2.38 cm-1, and |E/D| = 0.13. Please note that the negative value of D was confirmed by the temperature dependence of the signal (see supporting information, figs. S6 – S8). These values are typical for Mn(III) in proteins and compare well to those found in the photoexcited Mn(III) intermediate in photosystem II.40 The observed hyperfine couplings are similar to mononuclear Mn(III) model complexes.27, 28, 43, 46 The negative D together with the small hyperfine coupling constant suggest a 5B1 ground state which is expected either for a fivecoordinate square-pyramidal or a six-coordinate tetragonally elongated octahedral geometry.27 Zhu et al. reported fine structure and hyperfine structure values for Mn(III) (D = -4.0 cm-1, |E/D|


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= 0.11, A = 150 MHz) in citrate buffered OxDC under turnover conditions that are somewhat larger in magnitude than the ones we find here for the oxidized enzyme. As we will show below their values are indicative of the presence of the product molecule formate coordinated to the Nterminal Mn(III) ion.

Quantitative Analysis of Mn(III) in Untreated OxDC. It should not escape the attention of the reader that the untreated WT OxDC sample shows a weak Mn(III) signal in parallel mode as well (see fig. 1B, blue trace between 600 and 1000 G). It shows the same gvalue and hyperfine splittings as in the oxidized sample and is superposed by a large background signal. This observation provides clear evidence for the presence of Mn(III) in WT OxDC without the need for prior turnover or chemical oxidation and allowed us to quantitate the concentration of Mn(III) in virgin enzyme and compare it to earlier estimates by the Bornemann group.10, 13 Absolute spin quantitation is fraught with substantial error, particularly for parallel mode Mn(III) signals since they depend sensitively on the fine structure parameters, in particular the size of E. To minimize quantitation errors we elected to use relative intensity information in the untreated and oxidized spectra in perpendicular and parallel mode to derive an estimate of the Mn(III) concentration relative to the total EPR-visible Mn content. For this purpose, the blue and red traces in fig. 1, were replotted and intensities taken as the minimum-to-maximum intensity differences for the four low field lines of the parallel mode Mn(III) signal (see fig. 4B). The remaining two lines at higher field were considered unreliable because of potential interference with the parallel mode Mn(II) signal near geff ≈ 5.2. For quantitation we take the peak-to-trough intensity difference of a given line as the measure of its intensity and call it ci (i = 1, 2, 3, 4) for the oxidized sample and di for the untreated one. The ratio di/ci is calculated for each line position and then averaged over all four lines to give the fraction of the Mn(III) present in


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untreated sample compared to the oxidized one. The rationale for this approach is that averaging over four different line positions will minimize errors due to the unknown shape of the underlying baseline. This leads to a value of approximately 22%, i.e., the Mn(III) signal in the virgin enzyme sample has approx. 22% of the intensity of the Mn(III) signal in the oxidized sample. If all Mn(II) had been oxidized after treatment with iridate this number would mean that 22% of the EPR visible Mn was in the +3 oxidation state in virgin OxDC prepared in succinate buffer. However, we need to allow for a correction taking into account possibly incomplete oxidation in the iridate-treated sample. To this end a similar approach was taken to determine the relative intensities of the Mn(II) signals in the untreated, oxidized, and reduced samples while making allowance for dilution effects due to addition of small aliquots of stock iridate and ascorbate solutions during the course of the experiment. To estimate the fraction of Mn(II) that had been oxidized by the iridate treatment, three amplitudes were read from the perpendicular mode Mn(II) spectra near g ≈ 2, labeled ai and bi (i = 1, 2, 3) for the untreated and oxidized sample, respectively (see fig 4A). We only took the two low-field lines and the highest field line of the sextet into account as a measure of signal intensity because our cryostat has a weak but persistent copper background signal that is difficult to completely subtract out and may interfere with the central part of the Mn(II) spectrum. These signals in the g ≈ 2 region represent primarily the N-terminal Mn(II) site since the C-terminal site has significantly larger |D| values and therefore shows a much more spread-out and distorted spectrum.33, 38

Our quantitative analysis (see supporting information for details) shows that approximately 18% of the original g ≈ 2 signal is left over after oxidation representing the fraction / (see supporting information, table S1). This corresponds to about 7% of the total Mn content of the sample (see supporting information, table S2). A similar comparison may


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be performed on the half-field line in the perpendicular-mode EPR spectrum at g ≈ 4.2 which is a signature for the C-terminal Mn(II) (see supporting information, fig. S10). We find that approximately 29% of this signal is left over after oxidation is carried out, corresponding to approximately 16% of the total Mn content. This result suggests that the C-terminal Mn(II) is more difficult to oxidize than the N-terminal one. The same conclusion can be drawn from the high-field spectra of oxidized enzyme (vide infra) and indicates that the two Mn sites have slightly different redox potentials favoring the N-terminal Mn(II) as the one that more easily gives up an electron and may more readily serve as the catalytic site. Considering both the Nand the C-terminal Mn sites together we estimate that approximately 22-23% of the total Mn(II) content is left in the +2 oxidation state after the sample is oxidized with 8 mM hexachloroiridate with about twice as much C-terminal Mn(II) compared to N-terminal Mn(II). The remainder, approx. 77-78% has been converted to Mn(III) which then allows us to obtain an upper limit of 16% for the total EPR-visible Mn in the +3 oxidation state in virgin enzyme. Please note that this assumes that the signals for Mn(III) in the C- and in the N-terminal site are the same. If this is not the case our estimate has to be revised downward of the 16%.

Figure 4: Relative spin quantitation for the Mn(II) signal in perpendicular mode (A, left panel) and the Mn(III) signal in parallel mode (B, right panel). The blue spectra are untreated WT


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OxDC and the red represent the oxidized sample. Spectra are from the experiment shown in fig. 1. High-Field EPR Experiments. One of the difficulties with EPR on OxDC is the wide spread of fine structure parameters of Mn(II), especially for the C-terminal site.33, 38 Larger |D| and |E| results in the distribution of intensity over much larger parts of the spectrum and in the case of |D| > υ (the microwave frequency) the loss of certain transitions from the spectrum altogether at X-band. This complicates simulation of Mn(II) spectra at X-band where primarily the species with smaller magnitude D-values are detected near geff ≈ 2. The sharp lines often observed near geff ≈ 4 in perpendicular mode are most likely due to a non-principal axis turning point of the C-terminal Mn(II) with its much larger D-value in its penta-coordinated state. Based on this assignment our quantitative analysis of the X-band spectra (see above) suggests a smaller drop in intensity for the C-terminal Mn(II) than for the N-terminal one. Since both Mn(II) sites are much better resolved using high field EPR38, 47 we repeated the redox cycling experiment at 14.5 T/406.4 GHz (see fig. 5).

Figure 5: Redox titration of WT OxDC in 50 mM succinate buffer at pH4.2 using hexachloroiridate and ascorbate as the oxidizing and reducing agents, respectively. The experiment was carried out with the same sample thawed and refrozen multiple times between


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addition of aliquots of oxidant/reductant. Microwave frequency 406.4 GHz, 50 kHz modulation frequency, 1 G modulation amplitude, 0.2 mT/s field sweep speed. Sample temperature was set to 20 K.

Fig. 5 shows the results of this high-field EPR experiment on a sample of WT OxDC initially at a concentration of 25 mg/mL and pH4.2 which underwent successive additions of oxidant followed by spectrum acquisition at 20 K. At the end of the experiment ascorbate was added in excess to recover the Mn(II) species. Starting out with untreated WT OxDC, one can clearly distinguish two different Mn(II) species. They are identified as the N- and C-terminal Mn(II), species A and M in the notation of Tabares et al.38 In their 285 GHz spectra the Cterminal site M was identified as the low field shoulder on the sextet of lines. It is somewhat better resolved here at 406.4 GHz. As oxidant is added, both signals decrease in intensity and split further apart. The species at lower field (the C-terminal site) appears to vanish more slowly compared to the higher field sextet (the N-terminal site) in agreement with our analysis of the Xband experiments (vide supra). This suggests that the C-terminal Mn has a slightly higher reduction potential than the N-terminal one. After the sample has been fully reduced using ascorbate, the two sextet signals reappear but are more separated than initially and the Nterminal site has become more intense than before. This may be due to the fact that up to ~16% of the Mn may be initially in the +3 oxidation state and upon reduction may contribute to the Nterminal signal. It is of course also possible that the oxidation/reduction cycle has led to conformational changes on the C-terminal Mn binding site that would potentially lead to smaller |D| values which could mask the true increase in the spectrum of the N-terminal Mn(II).


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A similar oxidation/reduction experiment was performed on WT OxDC at high pH8.0 (see supporting information fig. S9). There, only the N-terminal Mn(II) shows up in a narrow sweep around g ≈ 2 because the C-terminal Mn is primarily pentacoordinated and has a very large |D| value which broadens the sextet making it considerably harder to observe.38 Upon oxidation with hexachloroiridate, the N-terminal Mn(II) disappears just as in the low-pH case. However, in contrast to low pH a carbon-based radical is observed (see also fig. 1A). This is most likely the tyrosyl radical that was found previously as the product of a side reaction in experiments performed at pH values between 5 and 6.26 High field affords the opportunity to resolve the g-anisotropy of organic radicals that might not be separated in X-band experiments.48, 49

Fig. 6 shows the trace observed at 406.4 GHz and 14.5 T when the enzyme is fully oxidized. A

rhombic g-tensor yields a reasonable simulation of the spectrum with gxx = 2.00680, gyy = 2.00394, gzz = 2.00179. In addition, a Gaussian g-strain of 0.001 was assumed for the gxx component to arrive at its wider line width.

Figure 6: Blue Trace: High-Field EPR of a carbon-based radical generated in the high-pH (8.0) WT OxDC sample by treatment with hexachloroiridate. Instrumental parameters: 406.4 GHz microwave frequency, 50 kHz modulation frequency, 1 G modulation amplitude, 0.2 mT/s sweep rate, 20 K temperature. Red trace: Simulation using a rhombic g-tensor (for parameters see text).


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The simulation parameters compare well with the expected g-tensor for tyrosyl radicals which had been observed before in other systems.50, 51 Since tryptophanyl radicals show very similar EPR parameters as tyrosyl radicals, final proof for this identification would have to be obtained using labeled tyrosine in OxDC which is beyond the scope of this contribution. We also attempted to observe the Mn(III) signals in oxidized samples of WT OxDC for both high and low pH using high field EPR. However, the sensitivity of our instrument is apparently not sufficient to detect them. It is not uncommon, especially in biological systems, that Mn(III) is difficult to observe due to the large D-strain that often accompanies protein environments.52

pH Dependence of the Mn(III) EPR Intensity. In order to further investigate the pH dependence of the Mn(III) signal, we utilized an immobilization protocol for OxDC that has recently been described.32 Briefly, the His6 tag of the enzyme, used for purification, was linked to Zn-loaded IMAC (immobilized metal affinity chromatography) resin. The resin beads were loaded on a home-made Kel-F® column (4×5 mm ID×OD) with a 5 µm polypropylene particle filter sheet placed at the bottom of the column to retain the beads. The column can be loaded with enzyme to a concentration of approximately 20 mg/mL and can be repeatedly frozen and thawed and the buffer exchanged by flushing the column. Enzyme loss has been observed to be less than 50% over the course of 10 freeze-thaw-flush cycles. Loss of enzyme due to low pH is also limited to less than 50%. Enzyme activity is lowered to approximately 25% by the immobilization procedure, presumably due to crowding effects on the resin beads. However, the EPR spectra of immobilized and free enzyme are very similar suggesting that the immobilization technique does not alter the protein conformation and the Mn binding sites. The loss of enzyme between experiment cycles prohibits absolute quantitation. However, it is possible to perform


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relative signal quantitation by comparing the intensities of the g ≈ 5.2 Mn(II) and the g ≈ 8.8 Mn(III) signals both observed in the same parallel mode spectra. Finally, in order to keep conditions as similar as possible during the variation of pH we employed a buffer cocktail using 50 mM each of Tris (pKa = 8.0), Bis-Tris (pKa = 6.5), piperazine (pKa = 9.4/4.9), succinate (pKa = 4.6/5.2), and 500 mM sodium chloride to span the pH range between 8.5 and 4.0. The pH was stepped down from the storage buffer pH of 8.5 initially to 8.0 and then in steps of one pH unit down to 4.0. With only five freeze-thaw-flush cycles we could be certain that most of the enzyme was retained on the column throughout the experiment and the same sample could be reused. Experimental spectra are shown in fig. 7A.

Figure 7: Panel (A) shows the parallel mode X-band spectra of immobilized WT OxDC at different pH. The experiment was started at pH8.5 and buffer was exchanged in discrete steps down to pH4.0. The sample was contained in a Kel-F® column that served also as the EPR sample tube for measurements at 5 K. The EPR parameters were: 9.384 GHz microwave frequency, 100 kHz modulation frequency, 10 G modulation amplitude, 0.63 mW microwave power. Panel (B) shows the intensity of the low-field line of the g ≈ 8.8 Mn(III) signal relative to (divided by the intensity of) the low field line of the g ≈ 5.2 Mn(II) signal as found in the same spectrum. Above pH7 no Mn(III) signal was observed.


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Upon lowering pH the geff ≈ 5.2 Mn(II) signal undergoes subtle changes in position and splitting as expected due to the pH dependence of the N-terminal Mn(II).38 However, at pH6.0 and below its shape stays the same indicating that the local geometry of the N-terminal Mn(II) site is not changing. Subtle absolute intensity changes are visible though and may in part be due to the partial loss of enzyme from the sample while flushing with buffer for the next pH step. Interestingly, the Mn(III) signal at geff ≈ 8.8 appears only at low pH. The relative intensity of the low-field lines of Mn(III) vs. Mn(II) is shown in fig. 7B and indicates a steady increase in Mn(III) concentration relative to the Mn(II) concentration as pH is lowered. This is reminiscent of the pH dependence of the logarithm of decarboxylase activity which follows a similar pH trend.12 Buffer Effects. Finally, we turn our attention back to the recent paper by Zhu et al.18 which reported parallel mode EPR of WT OxDC in citrate buffer at low pH. Interestingly, Mn(III) was not observed in their preparations until substrate was added. In fact the Mn(III) EPR signal did not even show up immediately after substrate addition as their rapid-freeze quench experiment testified, but only after thousands of turnovers (15 s after mixing whereas the turnover time is approx. 5 ms). The only difference between Zhu’s preparations and ours is the fact that they used citrate buffer while we typically use succinate buffer for low pH experiments. Turnover rates are similar for both preparations. Curious about this unexpected buffer effect we prepared enzyme in citrate buffer and indeed could not observe the Mn(III) signal in as-prepared samples (see fig. 8). We repeated Zhu’s experiments (see supporting information fig. S13) and confirm that Mn(III) may be observed in this preparation only after addition of substrate. In order to verify that it is the buffer that triggers the observation of Mn(III) in the absence of substrate we added a small aliquot of 1 M succinate buffer pH4.5 to the citrate-buffered sample for a final


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concentration of 100 mM and repeated the EPR measurement. The result is shown in fig. 8 (red trace).

Figure 8: X-band parallel mode EPR of WT OxDC at pH4.5 at 5 K. Black trace: sample in citrate buffer pH4.5, red trace: after addition of 100 mM succinate pH4.5, green trace: after addition of 100 mM acetate pH4.5 to the original citrate-buffered sample, blue trace: after addition of 100 mM formate pH4.5 to the original citrate-buffered sample.

The black trace shows WT OxDC in citrate buffer while the red trace is the same sample after addition of succinate buffer pH4.5 to a 100 mM final concentration. Only after succinate has been added the typical Mn(III) six-line spectrum near geff ≈ 8.8 appears near 800 G. The sextet splitting is identical to the one observed in succinate buffered OxDC at pH4.5 (see fig. 1B). It should be noted that the Mn(II) signal near geff ≈ 5.2 remains approximately the same upon addition of succinate (see supplementary information figure S14 where the same spectra are shown on a wider sweep range). A similar effect was observed when acetate or formate were added to a citrate buffered sample (green and blue trace in fig. 8, respectively). Clearly, the choice of buffer is important for the observation of Mn(III) in the absence of substrate. Citric acid has three carboxylic acid groups and is bulkier than succinic, acetic, and formic acid due to


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its branched structure. It is likely unable to pass through the solvent channel to coordinate to the N-terminal Mn, leaving it with two free coordination sites containing water. On the other hand, the other tested carboxylic acids are narrower in shape and may bind to the N-terminal Mn headon in a similar fashion as was recently demonstrated in a crystal structure containing oxalate bound to a Co ion substituted into the N-terminal binding site.18 It is well documented that complexation of metal ions affects their reduction potentials.53 This effect explains why the potential of the Mn(II)/Mn(III) couple is much lower in proteins than in aqueous solution where it is approx. 1.5 V.54 One can envision that the additional ligation with a negatively charged carboxylate group will further stabilize the Mn(III) in the N-terminal site making the corresponding Mn(II) easier to oxidize.

Table 1: Magnetic Parameters for Mn(III) simulated for OxDC in different buffers at low pH. Please note that the last column shows the values obtained for two different preparations, OxDC in citrate buffer pH4.5 in presence of formate and OxDC in citrate buffer with substrate, oxalate, added, which produces formate after turnover. The values for these two preparations are identical with each other and with the ones reported by Zhu et al.18 Succinate

Acetate

Formate/Oxalate

D [cm−1]

-2.38

-2.44

-4.0

|E/D|

0.13

0.13

0.11

A [MHz]

140

145

150

Our interpretation is qualitatively supported by the observation that the magnetic parameters (both fine structure and hyperfine structure) of the observed Mn(III) species is slightly different


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for the three different coordinating carboxylate species. The parameters giving the best fits to simulations (see supporting information, figs. S15 – S16) are shown in table 1.

It is significant that the simulation of the formate treated sample shows the same magnetic parameters as those observed by Zhu et al.18 Together with the fact that these workers only observed Mn(III) after thousands of turnovers proves that they did not observe a catalytically competent intermediate state but rather the product bound N-terminal Mn(III). The formate bound Mn(III) shows the largest negative D-value and the largest hyperfine coupling while the magnitude of D becomes smaller with increasing size of the coordinating carboxylate which may possibly be explained by an effect of decreasing charge density of the coordinating ligand on fine structure and hyperfine structure.


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Discussion The Mn(II) centers in OxDC have been well studied by EPR.26, 33, 38, 47 Despite their similar coordination to three histidine and one glutamate residues in both cupin domains the Nand C-terminal Mn(II) ions can be readily distinguished by their fine structure and its pH dependence.33, 38 Yet, mechanistic schemes in the literature typically invoke Mn(III) as the catalytically active redox state facilitating the initial 1-electron oxidation of oxalate10, 12-14, 19-21, 23, 26, 33, 34, 38, 47

(see scheme 1) similar to proposals for the mechanism of oxalate oxidase.25, 55-58 In

this scheme and related ones discussed in the literature it is always assumed that dioxygen, which acts as an obligatory co-catalyst, is bound to the active-site Mn center oxidizing it to form a catalytically competent Mn(III) redox state. There are three problems with this proposal that have rarely been discussed in the existing literature: (1) In order for dissolved dioxygen to generate Mn(III) the mismatch between the Mn(II)/Mn(III) and the O2/ O2·− potentials has to be overcome by the protein; (2) there is little direct spectroscopic evidence for Mn(III) (up till recently) or Mn(III)-O2·− in as-prepared WT OxDC; and (3) the close proximity of the O2·− and CO2·− radical intermediates in the same active site should lead to a strong propensity for oxidase activity. The recent publication by Zhu et al.18 attempted to address some of these questions and suggested that in order to form Mn(III) in OxDC, oxalate binds before dioxygen does. Moreover, the inability to detect a characteristic g ≈ 8.8 parallel mode signal which is specific for Mn(III) in certain coordination environments was taken as evidence that Mn(III) is absent in the resting state of the enzyme contrary to earlier reports by Bornemann’s group.10, 13 Our experiments show that the g ≈ 8.8 signal can indeed be observed in the low pH resting state of the enzyme in the presence of small molecules with carboxylate groups, specifically, formate, acetate, and succinate, while it is not observed in the presence of citrate. Quantitative analysis shows that up


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to 16% of the EPR-visible Mn is in the +3 oxidation state in the low pH resting state in very good agreement with Bornemann’s earlier results who also used succinate buffer. Analysis of the fine-structure of the Mn(III) signals shows characteristic differences for the three carboxylatecontaining molecules. The values for D and |E/D| for the three cases are listed in table 1. Comparison with Zhu et al.’s values reveals that these workers actually observed Mn(III) coordinated to formate. This makes perfect sense since they observed a relatively slow build-up of the Mn(III) signal commensurate with the formation of product.18 Specifically, their supporting information notes that Mn(III) could not be observed by rapid freeze quench at very short times after mixing oxalate with OxDC. The fact that formate, acetate, and succinate modulate the EPR parameters of Mn(III), specifically its D-value and its hyperfine coupling constant A, strongly suggests that these small molecules are binding to Mn(III) by inserting themselves into the N-terminal solvent channel. A simple explanation for the apparent absence of Mn(III) in the low pH resting state of OxDC in citrate buffer is that citrate as a branched molecule is too bulky to enter the active site while formate, acetate, and succinate, being all linear may end up coordinating with the Mn ion. The extra negative charge may be enough to stabilize Mn(III) over Mn(II) in the presence of dioxygen, i.e., it may lower the reduction potential of the Mn(II)/Mn(III) couple to be equal or lower than that of the O2/O2·− couple. It should be noted that this is more likely at low pH since the potential of the O2/HO2· couple is several 100 mV higher than that of O2/O2·−.15 In other words, the N-terminal Mn is poised at a reduction potential just above that of the O2/HO2· couple by the protein. Binding of substrate or another small carboxylate anion lowers its potential further and allows electron transfer from Mn(II) to O2 to happen. This initiates the catalytic cycle of oxalate dissociation or leads to an EPR-observable Mn(III) species when non-substrate carboxylate anions are coordinated.


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Whether dioxygen binds before or after the substrate cannot be decided by our experiments. While this is a reasonable hypothesis supported by our data, a word of caution is in order. The g ≈ 8.8 signal provides clear evidence of the presence of a mononuclear Mn(III) in the 5B1 ground state as noted before.27 However, its absence does not prove the absence of Mn(III) in the citrate-buffered system. In other words, if citrate can’t slip into the solvent channel and coordinate the N-terminal Mn might still be in the +3 oxidation state but in a different coordination environment with a possibly different ground state. Observation of such a species by X-band or even by high field EPR might be confounded if large strain and/or a very large or very small E parameter are present. The pH dependence of the EPR signal intensity of Mn(III) (see fig. 7B) bears a striking resemblance with the pH dependence of the enzymatic activity. This suggests that catalytic activity may not or not only depend on the presence of the oxalate monoanion which has been suggested as the catalytically active substrate based on its pKa of 4.2. Rather it seems likely that catalysis is governed by small pH-driven changes in the reduction potential of the O2/O2·− couple which shifts higher with lower pH and the presence of a negatively charged carboxylate group near the N-terminal Mn center which may stabilize its +3 oxidation state. The substrate oxalate would obviously be able to serve as such a coordinating carboxylate group and its binding to the N-terminal Mn(II) may actually trigger the initial electron transfer event that leads to formation of superoxide and Mn(III) which then initiates the catalytic cycle.


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Acknowledgement A plasmid containing the gene encoding C-terminally His6-tagged wild-type B. subtilis OxDC was originally provided by Dr. Stephen Bornemann (John Innes Centre, Norwich, UK). Useful technical discussions with Dr. Nigel G. J. Richards (University of Cardiff, School of Chemistry) are gratefully acknowledged.

Supporting Information Available Supporting information is available at: http://pubs.acs.org. Low pH step-wise redox cycling monitored by EPR, low pH step-wise redox cycling monitored by parallel mode EPR, high pH step-wise redox cycling monitored by EPR, high pH step-wise redox cycling monitored by parallel mode EPR, parallel mode EPR background signal, Mn(III) temperature dependence, Mn(III) temperature dependence simulation, high-field EPR high pH step-wise redox cycling, relative spin quantitation and quantitative analysi, parallel mode EPR under turnover conditions, Mn(III) buffer dependence, simulation of Mn(III) in the presence of acetate, simulation of Mn(III) in the presence of formate.


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AUTHOR INFORMATION Corresponding Author *Email: [email protected]. Present Addresses †Umar T. Twahir, Department of Physics, Emory University, 400 Dowman Drive, Atlanta, GA 30322, USA

Funding Sources Funding for this work was provided by the National Science Foundation under Grant CHE1213440. Part of this work was conducted at the NHMFL, which is funded by the NSF through a Cooperative Agreement DMR 1157490, the State of Florida, and the US Department of Energy.

Notes The authors declare no competing interest.


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in Oxalate Decarboxylase from Bacillus subtilis - ACS Publications

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