Elucidating the Role of O2 Uncoupling in the Oxidative Biodegradation of Organic Contaminants by Rieske Non-heme Iron Dioxygenases

Oxygenations of aromatic soil and water contaminants with molecular O2 catalyzed by Rieske dioxygenases are frequent initial steps of biodegradation in natural and engineered environments. Many of these non-heme ferrous iron enzymes are known to be involved in contaminant metabolism, but the understanding of enzyme–substrate interactions that lead to successful biodegradation is still elusive. Here, we studied the mechanisms of O2 activation and substrate hydroxylation of two nitroarene dioxygenases to evaluate enzyme- and substrate-specific factors that determine the efficiency of oxygenated product formation. Experiments in enzyme assays of 2-nitrotoluene dioxygenase (2NTDO) and nitrobenzene dioxygenase (NBDO) with methyl-, fluoro-, chloro-, and hydroxy-substituted nitroaromatic substrates reveal that typically 20–100% of the enzyme’s activity involves unproductive paths of O2 activation with generation of reactive oxygen species through so-called O2 uncoupling. The 18O and 13C kinetic isotope effects of O2 activation and nitroaromatic substrate hydroxylation, respectively, suggest that O2 uncoupling occurs after generation of FeIII-(hydro)peroxo species in the catalytic cycle. While 2NTDO hydroxylates ortho-substituted nitroaromatic substrates more efficiently, NBDO favors meta-substituted, presumably due to distinct active site residues of the two enzymes. Our data implies, however, that the O2 uncoupling and hydroxylation activity cannot be assessed from simple structure–reactivity relationships. By quantifying O2 uncoupling by Rieske dioxygenases, our work provides a mechanistic link between contaminant biodegradation, the generation of reactive oxygen species, and possible adaptation strategies of microorganisms to the exposure of new contaminants.


S2 Purification of nitroarene dioxygenases
Bacterial growth conditions and purification procedures were largely adapted from previous works. [2][3][4] In the following, we describe the complete purification protocol for the four enzyme components reductase, ferredoxin, 2-nitrotoluene dioxygense (2NTDO) and nitrobenzene dioxygenase (NBDO) adapted for this study. Chromatographic separations were performed on an automated fast protein liquid chromatography system (ÄKTA FPLC, GE Healthcare Life Science) with columns manually packed with resins from Cytiva (Marlborough, USA).

S2.2 Purification of ferredoxin
90 g of frozen cell pellet in BTGED buffer were thawed on ice and mixed with 1 mg mL −1 DNase I. The cell suspension was homogenised and passed twice through a chilled French pressure cell (1000 bar) and centrifuged at 10'000 g for 60 min at 4°C. The supernatant was passed through a 0.45 µm hydrophilic PVDF syringe filter and dialyzed in 2 L of fresh BTGED buffer and loaded onto a Q Sepharose XL ion exchange column (approximate bed volume of 200 mL) equilibrated with 600 mL BTGED. Unbound protein was removed with 200 mL BTGED at 2.5 mL min −1 followed by a linear gradient from 0 to 500 mM KCl in BTGED for 3 column volumes (CV). Fractions containing ferredoxin were identified by absoption at 456 nm 2 towards the end of the gradient and were combined and concentrated by ultrafiltration (10 kDa membrane, 1 bar nitrogen gas). Afterwards, ammonium sulfate was added to the concentrate to a final concentration of 1.5 M and the solution was loaded onto an octyl-sepharose 4 fast flow column (20 mL bed volume) equilibrated with 100 mL of 1.5 M (NH 4 ) 2 SO 4 in BTGED. Unbound protein including ferredoxin was removed with 50 mL of 1.5 M (NH 4 ) 2 SO 4 in BTGED at 1 mL min −1 . Fractions containing ferredoxin were pooled and concentrated by ultrafiltration. The concentrate was loaded onto a phenyl-sepharose 6 fast flow (high sub) column (20 mL bed volume) equilibrated with 100 mL 1.5 M (NH 4 ) 2 SO 4 in BTGED. Again, ferredoxin eluted with the unbound protein which was removed with 200 mL of 1.5 M (NH 4 ) 2 SO 4 in BTGED at 1 mL min −1 . Fractions containing ferredoxin were combined and concentrated by ultrafiltration. The buffer was exchanged to 50 mM MES (pH 6.8) and aliquots were stored at -80°C until needed. Table S1 gives an overview of the purification steps. 6

S2.3 Purification of reductase
74 g of frozen cell pellet in BTGED buffer were thawed on ice and mixed with 1 mg mL −1 DNase I. The cell suspension was homogenised and passed twice through a chilled French pressure cell (1000 bar) and centrifuged at 10'000 g for 60 min at 4°C. The supernatant was passed through a 0.45 µm hydrophilic PVDF syringe filter and dialyzed in 2 L of fresh  BTGED buffer and loaded onto a Q Sepharose XL ion exchange column (approximate bed volume of 200 mL) equilibrated with 600 mL BTGED. Unbound protein was removed with 200 mL BTGED at 2.5 mL min −1 followed by a linear gradient from 0 to 500 mM KCl in BTGED for 3 CV. Fractions containing recuctase were identified by absoption at 460 nm and activity tests 2 in the middle of the gradient and were combined and concentrated by ultrafiltration (30 kDa membrane, 1 bar nitrogen gas). Afterwards, ammonium sulfate was added to the concentrate to a final concentration of 1 M and the solution was loaded onto phenyl-sepharose 6 fast flow (high sub) column (20 mL bed volume) equilibrated with 100 mL 1.5 M (NH 4 ) 2 SO 4 in BTGED. Unbound protein was removed with 200 mL of 1 M (NH 4 ) 2 SO 4 in BTGED at 1 mL min −1 followed by a 12 CV gradient to 0 mM (NH 4 ) 2 SO 4 in BTGED. Fractions containing reductase eluted towards the end of the gradient and were pooled and concentrated by ultrafiltration. In contrast to previous procedures 2-4 , the third chromatographic step was replaced by size exclusion chromatography (SEC) due to an O 2 consuming impurity (see section S3.1.1). Pooled sample of less than 3 mL was loaded onto a HiLoad Superdex 200 pg (120 mL bed volume, GE Healthcare Life Science) equilibrated with 200 mL of MES buffer (50 mM, pH6.8) and run isocratically at 1 mL min −1 . Fractions containing reductase were combined and concentrated by ultrafiltration and aliquots were stored at -80°C until needed. Table S1 gives an overview of the purification steps. 6

S2.4 Purification of 2NTDO and NBDO
We performed identical steps for the purification of 2NTDO and NBDO as follows. 68 g of frozen cell pellet in BTGED buffer were thawed on ice and mixed with 1 mg mL −1 DNase I. The cell suspension was homogenised and passed twice through a chilled French pressure cell (1000 bar) and centrifuged at 10'000 g for 60 min at 4°C. The supernatant was passed through a 0.45 µm hydrophilic PVDF syringe filter and dialyzed in 2 L of fresh BTGED buffer and loaded onto a Q Sepharose XL ion exchange column (approximate bed volume of 200 mL) equilibrated with 600 mL BTGED. Unbound protein was removed with 200 mL BTGED at 2.5 mL min −1 followed by a linear gradient from 0 to 500 mM KCl in BTGED for 3 CV. Fractions containing oxygenase were identified by activity tests 2 in the beginning of the gradient and were combined and concentrated by ultrafiltration (100 kDa membrane, 1 bar nitrogen gas). Afterwards, ammonium sulfate was added to the concentrate to a final concentration of 1 M and the solution was loaded onto phenylsepharose 6 fast flow (high sub) column (20 mL bed volume) equilibrated with 100 mL 1.5 M (NH 4 ) 2 SO 4 in BTGED. Unbound protein was removed with 200 mL of 1 M (NH 4 ) 2 SO 4 in BTGED at 1 mL min −1 followed by a 12 CV gradient to 0 mM (NH 4 ) 2 SO 4 in BTGED.
Fractions containing oxygenase eluted after the end of the gradient and were pooled and concentrated by ultrafiltration. The buffer was exchanged to 50 mM MES (pH 6.8) and aliquots were stored at -80°C until needed. Table S1 gives an overview of the purification steps. 6

S3.1.1 Control experiments
We characterized the O 2 background consumption systematically with a number of control experiments shown in Table S2. The assays were set up as controlled substrate turnover experiments (see main manuscript) with initial NADH concentrations of 250 µM. Figure  S1 displays the O 2 consumption in the first 10 mins after NADH addition from which initial zero-order rates of O 2 consumption, ν 0 , were determined in µM min −1 .
Assays without any of the three enzymes, reductase (red), ferredoxin (fer), and the oxygenase of 2NTDO (oxy), or NADH did not consume any O 2 (|ν 0 | < 1). Assay without substrate consumed O 2 at a small rate of 2.92 ± 0.09 µM min −1 (Fig. S1a). This rate is still significantly smaller than the target reaction rate in the enzyme assay (17.9 ± 0.4 µM min −1 ). Controls with individual enzyme components (reductase, ferredoxin, and oxygenase) consumed less O 2 (Fig. S1b) suggesting that the O 2 consumption in the assay lacking substrate is not the result of an impurity but a reaction catalyzed by the full 2NTDO enzyme system. This is supported by the fact that no O 2 was consumed in assays without NADH. The modification of the reductase purification discussed in section S2.2 to include size exclusion chromatography (SEC), reduced the initial O 2 consumption rate from 1.24 ± 0.06 to 0.22 ± 0.09 µM min −1 . However, the O 2 consumption process evident in the "reductase only" samples does not appear to be as relevant when the whole multicomponent enzyme system is present as the impackt of the SEC purification was negligible in the "no substrate" samples (Table S2, entries 4 and 5).

Table S2
Control experiments for 2NTDO-catalyzed reactions performed for the assessment of background O 2 consumption in controlled substrate turnover experiments with nitrobenzene (NB). a Concentrations of assay components are indicated. Initial rates of O 2 consumption, ν 0 , were derived from linear fits to the data points obtained within the first 2 minutes after addition of NADH.
We tested the consequences of background O 2 removal in the various control assays for assessing the 2NTDO-catalysed activation of O 2 by analysing the remaining fraction of O 2 for changes in 18 O/ 16 O ratios. Figure S2 illustrates that with exception of the 'no substrate' assay (entry 3 in Table S2), no change in δ 18 O of O 2 were observed. Oxygen isotope fractionation in the 'no substrate' assay occurred to an extent that is comparable to experiments, where enzymatic O 2 activation is triggered by the presence of a substrate. Because this background reaction occurred at a > 6-fold slower rate, we exclude this source of O isotope fractionation as contribution to our experiments. The observation that the O isotope fractionation in the 'no substrate' assays were of similar magnitude as expected in the presence of the substrate suggest that some kind of oxydizable contamination might have been present in this assay. Further confirmation for neglecting such background O 2 consumptions and O isotope fractionation is shown in the following subsection.

S3.1.2 Quantification of O 2 uncoupling and background consumption
The extent of O 2 uncoupling, f O 2 -uc , was calculated through linear regressions of eq. S1 which corresponds to eq. 2 in the main manuscript.  Table S2. Experiments were initiated by addition NADH or the start of the O 2 measurement in case of the sample without NADH. [  Figure S3a shows linear regressions of eq. S1 for the 2NTDO-catalyzed dioxygenation of 2-nitrotoluene (f O 2 -uc of 0.02±0.03) and 4-chloronitrobenzene (f O 2 -uc of 0.92±0.01). A slope close to unity, as for 2-nitrotoluene, implies a stoichiometric transfer of activated O 2 species to the substrate and formation of (substituted) catechols. By contrast, only about 8% of the activated O 2 is used for dioxygenation in reactions with 4-chloronitrobenzene and the remainder is released as reactive oxygen species. Figure S3b illustrates that, in some assays, the fit did not go through the origin (0|0) but had positive or negative intercepts, b, of −36 ± 2.8 to 8.22 ± 5.38 µM (Table S3). Positive intercepts are small and close to 0 within the margin of error and were not evaluated further. Negative intercepts, however, indicate a gap where a constant amount of dissolved O 2 was removed in all of the assays in one set of experiments, regardless of the NADH concentration. We interpret this gap as a O 2 background consumption, [O 2 ] bg , quantified with eq. S2.  are unclear and no link to purification batches was observed. Qualitatively, we observed a negative trend of [O 2 ] bg with increasing f O 2 -uc ( Figure S4a). We interpret this as an artifact of the evaluation method for f O 2 -uc (eq. S1) because measurement uncertainties are larger in the measurement of small quantities of NO -2 and nitrobenzylalcohol and even a small systematic overestimation leads to a positive intercept, b, resulting in high negative values of [O 2 ] bg . This is also supported by the fact that all negative values of [O 2 ] bg exhibit high 95% confidence intervals that make them indistinguishable from 0.
In the data evaluation, we took the phenomenon of O 2 background consumption, [O 2 ] bg , into account as follows. The good quality of the linear fit ( Figure S3a) is an indicator of the accuracy of the data. Inclusion of the NADH = 0 sample, where no O 2 is consumed and no NO -2 is formed, in the raw data as (0|0), however, would distort the linear fit. Instead, we corrected the O 2 consumption, ∆O 2 , by the value of [O 2 ] bg ( Figure S3b). As the background consumption of O 2 is not the result of dioxygenation reactions, we excluded positive values from the calculation of |υ O 2 | with eq. 2 in the main manuscript by correcting O 2 concentrations with eq. S3.  Figure S4b shows that the O 2 background consumption did not affect this value. We speculate that the unknown process leading to background consumption of O 2 in 2NTDO assays with selected substrates might exhibit a similar 18 O-KIE and we refrained modifying the quantification of 18 O-KIEs.

S3.2.2 Quantification of dissolved O 2
Concentrations of aqueous, dissolved O 2 were measured continuously with needle-type fiber-optic oxygen microsensors connected to a 4-channel transmitter (PreSens Precision Sensing GmbH) as reported previously. 7 Up to 4 oxygen sensors were operated simultaneously after daily calibration with air-saturated and oxygen-free water. O 2 concentrations were corrected for variations in temperature. The analytical uncertainties of O 2 concentrations were smaller than ±0.5 µM.

S3.3 Enzyme Kinetics
The kinetics of initial O 2 consumption and NO -2 formation in the presence of different substrates i were evaluated in separate assays (see main manuscript). For the sake of comparability, all kinetic data were determined with the same batch of purifications. Initial rates of nitrite formation, ν i 0,NO − 2 , were obtained from repeated sampling during the first 60 sec after substrate addition. Initial rates of O 2 consumption, ν i 0,O 2 , were determined from continuous measurements of dissolved O 2 concentration, c O 2 , during the first minute after NADH addition ( Figure S5).
Maximum rates (ν i max ) and Michaelis constants (K i m ) of nitrite formation in the presence of different substrates i were determined with a non-linear least square regression according to eq. S4, is the initial rate of NO -2 formation from substrate i, c i 0 is the nominal initial substrate concentration, k i cat is the observable first-order rate constant, and E 0 is the nominal concentration of active sites in NBDO, corresponding to 3 mol per mol of oxygenase. By contrast, ν i max and K i m for O 2 consumption were obtained from the continuous measurement of O 2 concentration (c O 2 ) over time in a single assay. The rate of O 2 consumption at each time-point (ν i O 2 ) was calculated as the derivative of measured c O 2 vs. time (i.e., ∆[O 2 ]/∆t). We used non-linear least square regression according to eq. S5 with the derived ν i O 2 and measured c i O 2 values was used to estimate ν i max and K i m .
S3.4 13 C/ 12 C ratio analysis in substrates with limited turnover In this study, the quantification of species concentration as well as stable isotope ratios of oxygen and carbon in O 2 and organic substrates were all performed from aqueous and gaseous samples from the identical reactors. To that end, the very inefficient oxygenation of some substrates by 2NTDO and the concomitant extensive O 2 consumption compromised the quantification of their C isotope fractionation and 13 C-KIE values. As shown in Table S4, the turnover of 4-nitrotoluene, as well as the three nitrophenol isomers in assays of 2NTDO, 1 − c/c 0 , was particularly low (i.e. < 0.4) or even neglible. The C isotope fractionation of these substrates was relatively minor and mostly below the total uncertainty of 0.5 for 13 C/ 12 C ratio measurements. 9 To that end, 13 C-KIE values of nitrophenol isomers were set to unity. For 4-nitrotoluene, we used the 13 C-KIE of 1.003 ± 0.001 determined in whole cell experiments with E. coli clones expressing 2NTDO. 10   Table S4 Maximum substrate consumption, 1−c/c 0 , of various substrates in assays of 2NTDO and observed 13 C/ 12 C fractionation as changes in δ 13 C.
[S] 0 is the nominal initial substrate concentration used.  Figure S5 and Table S5. We observed three distinct types of substrate-dependent behaviour. First, various well-known nitroaromatic substrates consume O 2 with concomitant generation of NO -2 from the dioxygenation reaction. Their O 2 consumption is shown in Figure S5a. These are summarized as type-1-substrates in Table S5 and include new substrates such as dichloronitrobenzene and nitronaphthalene. Second, we identified nitroaromatic substrates that cause O 2 disappearance without generation of NO -2 . These substrates, summarized as type-2-substrates in Table S5 include the explosives 2,4,6-trinitrotoluene and 2,4-dinitroanisole which, like nitrophenols, act as O 2 uncoupling compounds. Finally, we find that the O 2 consumption of aromatic compounds lacking a NO 2 -group do not consume any O 2 beyond background losses. These compounds include benzene, toluene, naphthalene, and benzoate and no efforts were made to measure any of the expected dioxygenation products.  Table S5).

S4.2 H 2 O 2 quantification
We quantified H 2 O 2 concentrations in enzyme assays based on horse radish peroxidase catalyzed turnover of either Ampliflu™ or p-methoxyanilin. Results are presented in Table S6.