Expression, purification and UV-visible absorption features of CYP116B1
The gene encoding the C. metallidurans CH34 CYP116B1 P450 (gene 4932, UniProt Q1LDI2) was cloned by PCR from bacterial genomic DNA and expressed in Escherichia coli (Origami (DE3)) using a pET11a construct. CYP116B1 was purified using a combination of ion-exchange and hydrophobic affinity chromatography, producing a single major band on SDS/PAGE (Fig. 2A, inset), consistent with the predicted 86.4-kDa molecular mass of the P450–PDOR fusion protein. The UV-visible absorption properties of the pure CYP116B1 in its oxidized, sodium dithionite-reduced and reduced/carbon monoxide (CO)-bound forms are illustrated in Fig. 2A. For oxidized CYP116B1, the spectral features of a haem cofactor are clearly evident. The main Soret band has its absorption maximum at 418 nm, with longer-wavelength alpha/beta bands positioned at 566 and 532 nm, respectively. A pyridine haemochromagen assay to determine the haem concentration of CYP116B1 was performed in triplicate and a CYP116B1-specific extinction coefficient was determined to be ε418 = 121 mm−1·cm−1 at the Soret peak of the oxidized enzyme. This value includes the spectral contributions from iron-sulfur and flavin centres in the cofactor-replete CYP116B1, and was used to determine the CYP116B1 concentration in all further experiments.
Figure 2. UV-visible absorption spectroscopy of CYP116B1. (A) UV-visible absorption spectra for CYP116B1 (2.9 μm) showing the purified oxidized enzyme (thick solid line) with Soret maximum at 418 nm, the dithionite-reduced enzyme (dashed line) and the reduced/CO-bound adduct (thin solid line) showing the Fe(II)CO peak at 449 nm with a minor peak at 420 nm from the (protonated) cysteine thiol-bound form. Inset: SDS/PAGE gel (10%) of CYP116B1 showing the purified enzyme at 86.4 kDa at three different concentrations of 1, 0.5 and 0.1 μg (left to right), sizes of marker bands indicated. (B) Complexes of CYP116B1 with small-molecule inhibitors. Spectra shown are for oxidized CYP116B1 (2.3 μm, thick solid line); NO-bound CYP116B1 (thin solid line), showing a Soret intensity decrease and shift to 433 nm with Q-bands at 574 and 545 nm; cyanide (CN)-bound (50 mm KCN) CYP116B1 (dotted line), showing a Soret intensity decrease and shift to 430 nm; and imidazole (2 mm)-bound CYP116B1 (dashed line), exhibiting a Soret shift to 424 nm with a small decrease in intensity. Inset: CYP116B1-imidazole binding curve generated from difference spectral data (as described in the Materials and methods), yielding a Kd value of 646 ± 13 μm.
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On aerobic reduction with sodium dithionite, the Soret band decreased in intensity and was slightly blue-shifted to 417 nm. The less intense haem alpha and beta bands remained at ∼ 566 and 532 nm, respectively. The extent of Soret absorbance shift was less than that observed for other P450s (e.g. the P450 BM3 Soret shifted to ∼ 409 nm on full reduction to the ferrous, thiolate-coordinated state) [35,36]. The less-extensive Soret shift for CYP116B1 is a consequence of only partial reduction of the haem iron by dithionite under aerobic conditions, which, in turn, is a result of the rapid reaction of its ferrous haem iron with dioxygen and its subsequent re-oxidation with production of superoxide. Anaerobic titration of CYP116B1 with dithionite led to complete reduction of the haem iron and a more extensive Soret shift (see the section ‘Examination of the thermodynamic properties of CYP116B1’). However, the simultaneous drop in absorption intensity in the region beyond ∼ 425 nm suggests that the major cause of the apparent Soret absorption decrease is actually the reduction of, and concomitant changes in the spectral contributions of, the other cofactors bound to the protein. The reductase (PDOR) domain (amino acids ∼ 445–780 of the 780-residue CYP116B1) is expected to bind both flavin (FMN) and iron–sulfur cofactors, by homology with both the B. cepacia PDOR and the reductase domain of Rhodococcus sp. NCIMB9784 CYP116B2 . Reductive bleaching of FMN, and 2Fe-2S cofactor absorption by dithionite thus underlies the decreased Soret intensity and the absorption bleaching observed beyond ∼ 425 nm.
In view of the differing coenzyme selectivity of the B. cepacia PDOR (which favours NADH) and the Rhodococcus CYP116B2 (which favours NADPH) [30,32], we also analyzed spectral changes on aerobic mixing of oxidized CYP116B1 with NADH and NADPH (50 μm). Interestingly, spectral changes similar to those observed with dithionite were seen using NADPH as reductant, but NADH only effected partial reduction of the FMN/iron–sulfur centres (data not shown). In view of our subsequent work (vide infra), which shows that both NADPH and NADH support CYP116B1 catalytic activity, it is likely that this reflects faster enzyme reoxidation (and/or slower reduction) for NADH-reduced CYP116B1 compared with the NADPH-reduced form. This conclusion is consistent with stopped-flow studies reported later, under ‘Steady-state and stopped-flow kinetic studies on CYP116B1’.
Addition of CO to the dithionite-reduced CYP116B1 resulted in a shift of the Soret peak to 449 nm, with a much smaller feature at ∼ 424 nm. The former band is the characteristic signature for the cysteine thiolate-ligated P450 haems in their Fe(II)CO complexes. The minor feature at ∼ 424 nm probably reflects a small proportion of the (protonated) cysteine thiol-coordinated P420 form. Protonation of the proximal ligand can arise as a consequence of either haem reduction or the binding of CO to ferrous haem iron, or both [37,38]. Other spectroscopic studies indicate that the cysteine thiolate-coordinated state is predominant in the oxidized enzyme (see ‘Characterization of CYP116B1 haem and iron–sulfur clusters by EPR’). In the Fe(II)CO complex of CYP116B1, there is a single absorption Q-band peak in the visible region at ∼ 549 nm, indicative of complete conversion of the haem iron to the ferrous state (Fig. 2A).
To further define the spectral features of CYP116B1, we generated ligand complexes using the small-molecule inhibitors imidazole, cyanide and nitric oxide (NO) (Fig. 2B). All three molecules induced red (type II) shifts in the Soret band, which is typical for haem iron-coordinating P450 inhibitors. The imidazole complex had its Soret band shifted to 423.5 nm with a small decrease in intensity. An optical binding titration with imidazole revealed a hyperbolic dependence of induced absorption change on ligand concentration and a Kd of 646 ± 13 μm (Fig. 2B, inset). The Soret shift from ∼ 418 nm to 423.5 nm on complexation of imidazole and triazole inhibitors is typical of behaviour seen in other P450s (e.g. 38,39). However, it was reported that purified Rhodococcus CYP116B2 has its Soret maximum at ∼ 424 nm in the ligand-free state . Given that this enzyme was purified by imidazole elution from nickel resin, it is probably the case that the CYP116B2 form purified was the imidazole complex. The CYP116B1 complex with (sodium) cyanide resulted in a Soret shift to 431 nm and a substantial decrease in peak intensity (as also observed, for example, in the Mycobacterium tuberculosis sterol demethylase CYP51B1) . Optical titration indicated a Kd value of 9.7 ± 0.6 mm for cyanide. Finally, binding of the gaseous ligand NO led to a Soret shift to 433 nm, with a distinctive sharpening of features in the visible region (Q-bands at 574 and 545 nm), and a spectrum comparable to the NO-bound form of P450 BM3 (Fig. 2B) .
The binding of P450 substrate molecules often induces spectral shifts in the Soret region. Blue shifts (towards ∼ 390 nm) are observed when substrates cause displacement of an aqua ligand bound weakly to the sixth coordination position on the haem iron, as seen, for example, for camphor binding to P450cam (CYP101A1) and for fatty acid binding to P450 BM3 [41,42]. To identify potential substrate molecules for CYP116B1, we undertook extensive binding trials with a diverse range of prototypical P450 substrates, including fatty acids (e.g. lauric acid and myristic acid), steroids and other cyclic molecules (e.g. testosterone, hydrocortisone, 7-ethoxycoumarin and 6-deoxyerythronolide B). While there were no significant optical changes observed with the saturated fatty acids tested (lauric acid [C12], myristic acid [C14], pentadecanoic acid [C15] and palmitic acid [C16]), small type-I spectral shifts (typically 2–3 nm, spin-state change of ∼ 4–10%) were seen with a number of monounsaturated fatty acids (myristoleic acid [C14], palmitoleic acid [C16] and oleic acid [C18]), and with the polyunsaturated C20 arachidonic acid. The low solubility of oleic acid in aqueous buffer did not allow for affinity to be determined by optical titration. However, this was feasible for the three other lipids, and data for induced optical change versus fatty acid concentration were fitted using a hyperbolic function to generate Kd values of 4.8 ± 1.7 μm (myristoleic acid), 33.5 ± 6.3 μm (palmitoleic acid) and 5.6 ± 1.3 μm (arachidonic acid). Data are shown for myristoleic acid in Fig. 3A. Given the successful demonstration of binding of fatty acid molecules to CYP116B1, we also investigated whether fatty acid-linked imidazole inhibitors (imidazolyl decanoic acid [ImC10], imidazolyl undecanoic acid [ImC11] and imidazolyl dodecanoic acid [ImC12]) could act as potent inhibitors of P450. In previous studies we demonstrated tight binding of ImC10, ImC11 and ImC12 to the P450 BM3 fatty acid oxygenase . These molecules also bound tightly to CYP116B1 and induced type-II spectral changes (Soret shifts to 424 nm), consistent with haem iron coordination. The Kd values obtained from fitting optical-titration data using a quadratic equation for tight-binding ligands were 3.5 ± 0.4 μm (ImC10), 0.59 ± 0.12 μm (ImC11) and 0.52 ± 0.11 μm (ImC12) . Data for the binding of ImC12 to CYP116B1 are shown in Fig. 3B. These Kd values compare favourably to those determined for binding of ImC10, ImC11 and ImC12 to P450 BM3 (0.9, 7.5 and 1.35 μm, respectively), and with data for binding to a rat CYP4A1–CPR fatty acid hydroxylase fusion protein (12.4, 0.37 and 0.35 μm, respectively) [43,45].
Figure 3. Substrate- and inhibitor-type binding in CYP116B1. (A) Binding curve for myristoleic acid-induced spectral change in CYP116B1 (2.0 μm). Data were fitted using a quadratic function , generating a Kd value of 4.8 ± 1.7 μm for myristoleic acid. Inset: difference spectra showing the progressive substrate-like Soret absorbance shifts to shorter wavelength (blue shifts) upon addition of myristoleic acid at 7.96, 15.92, 23.88, 47.76 and 79.60 μm. (B) Binding curve for ImC12-induced spectral change in CYP116B1. Data were fitted as described for (A), producing a Kd value of 0.52 ± 0.11 μm. Inset: UV-visible absorbance spectra for CYP116B1 in ligand-free form (2.1 μm, solid line) and with ImC12 at a near-saturating concentration (5.13 μm, dashed line). There is a shift in the Soret peak maximum from 418 to 424 nm on binding ImC12.
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In view of the close relationship (52% amino-acid sequence identity) of the CYP116B1 haem domain with the R. erythropolis CYP116A1 (also known as ThcB, a non-fused P450 shown to catalyse the N-dealkylation of EPTC ), we also analysed the binding of EPTC and the related thiocarbamate herbicide S-propyl dipropylthiocarbamate (vernolate) to CYP116B1. Very small type-I haem perturbations were observed, but these were not large enough to enable accurate determination of Kd from optical binding. However, we were able to show that CYP116B1 catalyses the oxidation of both vernolate and EPTC (see ‘Identification of CYP116B1-catalysed reaction products of thiocarbamate herbicides’).
Characterization of CYP116B1 haem and iron–sulfur clusters by EPR
EPR spectra for the oxidized and sodium dithionite-reduced forms of pure CYP116B1 were collected as described in the Materials and methods. For oxidized CYP116B1, a rhombic spectrum consistent with a ferric cysteinate-coordinated haem iron was observed, with g-values of 2.42 (gz), 2.25 (gy) and 1.93 (gx) (Fig. 5). These g-values lie in the range typical for other low-spin P450 enzymes [e.g. 38,47,48], and are consistent with an essentially completely low-spin P450 enzyme. There were no significant signals in the region associated with the high-spin P450 haem iron (e.g. g = 8.18/7.85, 3.44/3.97 and 1.66/1.78 for P450 BM3/P450cam in substrate-bound forms [48,49]), suggesting that ferric CYP116B1 is hexacoordinated with a water ligand trans to the cysteine thiolate. Similar sets of g-values were obtained for the low-spin forms of other bacterial P450s. These include P450 BM3 and P450 BioI from Bacillus subtilis (CYP107H1), whose EPR spectra yield g-values of 2.42, 2.26 and 1.92, and 2.41, 2.25 and 1.92, respectively [47,48].
Figure 5. X-Band EPR spectra for CYP116B1. Upper spectrum: oxidized CYP116B1 (184 μm) showing a rhombic signal with g-values typical for a low-spin, six-coordinate ferric haem iron at gz = 2.42, gy = 2.25 and gx = 1.93. Lower spectrum: CYP116B1 (184 μm), reduced anaerobically by dithionite, showing almost complete loss of the signal from the ferric haem species (as haem is reduced to the EPR-silent ferrous state) and the emergence of bands at gz = 2.06, gy = 1.97 and gx = 1.88, typical of a plant-like 2Fe-2S cluster in the EPR active [2Fe-2S]+ form. No significant signal is observed for a flavin semiquinone.
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For oxidized CYP116B1, there were no apparent EPR signals that could be ascribed to either iron–sulfur or flavin centres that are likely to be present in the PDOR domain of the enzyme. Oxidized flavins are EPR silent, and the 2Fe-2S cluster in its oxidized form (i.e. [2Fe-2S]2+) should contain two ferric iron atoms, in which the magnetic moments of the unpaired electrons are spin-coupled, giving S = 0. However, reduced 2Fe-2S clusters are EPR active in the [2Fe-2S]+ form, with one ferric and one ferrous iron atom [e.g. 50,51]. An EPR spectrum was collected for the dithionite-reduced form of CYP116B1 and is shown in Fig. 5. The spectrum is more compact than that for the oxidized enzyme with the major features appearing between 3250 and 3750 Gauss. The reduced CYP116B1 spectrum has a rhombic shape and g-values of gz = 2.06, gy = 1.97 and gx = 1.88 that fall within the typical ranges reported by More et al. , clearly identifying the presence of a reduced 2Fe-2S cluster. There is no EPR signal that could be clearly assigned to a flavin semiquinone in the reduced CYP116B1. The EPR spectrum for the 2Fe-2S cofactor-reconstituted PDOR (2Fe-2S- and FMN-binding) domain of CYP116B2 from Rhodococcus sp. NCIMB 9784 produces an identical set of g-values . The CYP116B1 2Fe-2S EPR spectrum is also highly similar to that for the homologous B. cepacia PDOR (g = 2.04, 1.95 and 1.90), and to that of the Arabidopsis photosynthetic ferredoxin Fd2 (g = 2.05, 1.95, 1.89) [53,54]. Integration of the EPR signals for the oxidized haem and for the reduced iron–sulfur cofactors in CYP116B1 indicate that the 2Fe-2S centre is present at ∼ 98% of the concentration of the haem, thus suggesting near-complete incorporation of the iron-sulfur cluster in CYP116B1.
To identify the flavin bound to CYP116B1, we isolated the bound cofactor from a purified enzyme sample, as described in the Materials and methods. Thereafter, flavin fluorescence was determined at both pH 7.7 and pH 2.0, as described previously , and compared with FAD and FMN standards treated in the same way. Decreased flavin fluorescence was observed for both the CYP116B1 cofactor extract and for FMN standards on acidification, while enhanced fluorescence was seen for FAD (owing to hydrolysis of its pyrophosphoryl bond and release of ADP and the more highly fluorescent FMN). FMN content was estimated at ∼55% by comparison with the quantity of bound haem, consistent with the previous report of variable FMN content in CYP116B2 . The inclusion of free FMN (1 mm) in buffers used for CYP116B1 purification enabled increased FMN content in the purified enzyme (typically to > 80%).
Steady-state and stopped-flow kinetic studies on CYP116B1
Analysis of the ability of CYP116B1 to reduce electron acceptor molecules (potassium ferricyanide, cytochrome c) in a NAD(P)H-dependent manner and to perform P450 substrate-dependent oxidation of NAD(P)H was examined in steady-state reactions (Table 1). There was a strong preference for NADPH over NADH, with a difference of ∼ 440-fold in Km values for NADPH (0.9 μm) and NADH (399.1 μm) determined in cytochrome c-reduction studies. The kcat values for both ferricyanide and cytochrome c reduction were also ∼three-fold greater with NADPH as the coenzyme than with NADH (e.g. 833.1 min−1 versus 286.8 min−1 for ferricyanide), resulting in a 99-fold difference in catalytic efficiency for ferricyanide reduction (measured as the ratio of kcat/Km[NADPH] to kcat/Km[NADH]) and a 1174-fold difference for cytochrome c reduction in favour of NADPH (Table 1). CYP116B2 also demonstrated preference for NADPH (Km = 3.0 μm) over NADH (102.1 μm) in steady-state ferricyanide-reduction assays . In contrast, the B. cepacia PDOR flavoprotein has strong preference for NADH (Km ∼ 10 μm) over NADPH, and NADPH did not support measurable activity with either cytochrome c or the partner protein PDO, pointing to an important evolutionary divergence between the stand-alone PDOR enzyme and the fused PDOR domains in the CYP116B enzymes.
Table 1. Steady-state kinetic properties of CYP116B1. Km and kcat values for CYP116B1 were determined from steady-state kinetic assays with NADH and NADPH as reductants, and using the electron acceptors potassium ferricyanide (FeCN) and cytochrome c, and the P450 substrates vernolate and EPTC. Km values shown are for NAD(P)H in the experiments using cytochrome c and FeCN, and for vernolate/EPTC in the experiments using the thiocarbamates. In experiments using the thiocarbamate substrates, NADH (1.5 mm) and NADPH (200 μm) were used at near-saturating concentrations. All assays were performed on a dual-beam spectrophotometer (Jasco) in buffer A at 25 °C using CYP116B1 at a final concentration of 75–100 nm
|Acceptor or substrate||Reductant|
| Km (μm)||0.9 ± 0.5||399.1 ± 52.1|
| kcat (min−1)||151.1 ± 8.0||57.1 ± 2.7|
| Km (μm)||3.0 ± 1.0||102.1 ± 12.2|
| kcat (min−1)||833.1 ± 17.8||286.8 ± 7.5|
| Km (μm)||210.2 ± 30.7||270.4 ± 58.8|
| kcat (min−1)||57.3 ± 1.9||34.0 ± 2.2|
| Km (μm)||50.3 ± 9.6||110.4 ± 10.0|
| kcat (min−1)||30.7 ± 0.5||17.0 ± 0.4|
In view of the preceding data indicating substrate-like (type I) Soret spectral shifts induced on binding both unsaturated fatty acids and (to a lesser extent) the thiocarbamate herbicides EPTC and vernolate, we analysed steady-state oxidation of NADPH by CYP116B1 in the presence of these potential substrates. Despite favourable haem optical changes induced by myristoleic, palmitoleic, oleic and arachidonic acids, none of these molecules stimulated CYP116B1-dependent oxidation of either NADH/NADPH to any significant levels. By contrast, substantial stimulation of oxidation of both coenzymes was observed with EPTC and vernolate as substrates. A hyperbolic dependence of coenzyme oxidation rate on EPTC concentration was obtained using both NADH and NADPH, and data were fitted using the Michaelis function to produce values of kcat = 30.7 min−1 and Km EPTC = 50.3 μm (with NADPH as the donor) and kcat = 17.0 min−1 and Km EPTC = 110.4 μm (with NADH as the donor). For vernolate, the comparable values were kcat = 57.3 min−1 and Km vernolate = 210.2 μm (using NADPH as the donor) and kcat = 34.0 min−1 and Km vernolate = 270.4 μm (using NADH as the donor) (Table 1). Thus, EPTC (a substrate that is dealkylated in whole cells of R. erythropolis that express CYP116A1) clearly stimulates NAD(P)H oxidation in CYP116B1, and the related dithiocarbamate vernolate enhances NAD(P)H oxidation to an even greater extent. In the absence of vernolate, NADPH-dependent reduction of the CYP116B1 haem iron is slow in a CO-saturated solution (∼ 0.2 min−1) and results mainly in the P420 form of the enzyme. In the presence of saturating vernolate, there is some stimulation of the rate of haem iron reduction, but data analysis is complicated by a greater tendency of the enzyme to aggregate with development of turbidity in the solution. Despite the progressive aggregation of vernolate-bound CYP116B1 in this experiment, it is notable that the predominant species is now the P450 form, indicating substrate-dependent stabilization of the thiolate ligand, as observed previously in other systems [37,38,56]. Given the limited solubility of vernolate, its marginal effect on haem iron spin-state equilibrium and its influence on aggregation of CYP116B1, it is difficult to establish the extent to which electron transfer to the CYP116B1 haem iron is thermodynamically gated in the presence of this substrate and a kinetic gate to haem reduction in CYP116B1 cannot be ruled out . Consistent with the monomeric form of CYP116B1 being the catalytically relevant species, specific rates of vernolate-dependent NADPH oxidation, determined across a range of CYP116B1 concentrations (10–500 nm), showed no significant variation. In contrast, similar studies with P450 BM3 and the substrate lauric acid showed diminished specific activity at low enzyme concentration as a result of disassociation of the catalytically active dimer .
To further characterize the reaction of CYP116B1 with NAD(P)H, stopped-flow absorption spectroscopy was used to analyse the kinetics of reduction of CYP116B1 by NADH and NADPH. Absorption transients were collected at 465 nm (i.e. near the expected absorption maximum for the oxidized FMN cofactor ) after mixing CYP116B1 with different concentrations of NAD(P)H. Reaction transients were biphasic in all cases and were fitted accurately using a double exponential function. Approximately 80% (NADH) and 90% (NADPH) of the overall absorption change observed on coenzyme-dependent reduction of the FMN/2Fe-2S centres occurred in the fast phase of the reaction transients. The CYP116B1 haem was not reduced by NAD(P)H to any extent in its substrate-free form. Figure 6A shows a plot of the individual rate constants for the fast (kobs1) and slow (kobs2) phases of the transients against NADH concentration, while Fig. 6B shows the same data set with NADPH as reductant. For NADH-dependent reduction, kobs1 shows a hyperbolic dependence on the NADH concentration, and data were fitted to yield a limiting rate constant (klim) of 21.9 ± 0.2 s−1 and an apparent NADH Kd of 21.7 ± 0.8 μm. The kobs2 values for NADH are much slower (0.47 ± 0.03 s−1) and show no dependence on the NADH concentration (Fig. 6A). For NADPH-dependent reduction of CYP116B1, the reaction rate constants are higher, but there is no dependence of either kobs1 (72 ± 6 s−1) or kobs2 (5.5 ± 3.0 s−1) on the NADPH concentration (Fig. 6B).
Figure 6. Stopped-flow kinetic analysis of NADH- and NADPH-dependent CYP116B1 reduction. Stopped-flow absorption kinetic studies of CYP1161B1 (3.3 μm final concentration) were performed by monitoring the reduction of FMN/2Fe-2S centres at 465 nm (the wavelength of maximal overall change between oxidized and reduced CYP116B1) using both NADH (A) and NADPH (B) reducing coenzymes at a range of concentrations up to 500 μm. Open circles represent individual rate constants derived from the fast phase (kobs1) of double-exponential fits to the ΔA465 versus time transients, while open squares represent kobs2 rate constants for the slow phase of the reaction transients. There is no apparent dependence of either kobs1 (72 ± 6 s−1) or kobs2 (5.5 ± 3.0 s−1) on [NADPH]. With NADH as reductant, there is a hyperbolic dependence of kobs1 on [NADH], giving a limiting rate constant (klim) of 21.9 ± 0.2 s−1. However, there is no apparent dependence of kobs2 on [NADH] (0.47 ± 0.03 s−1).
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Examination of the thermodynamic properties of CYP116B1
To analyse the redox properties of CYP116B1, a spectroelectrochemical redox titration was performed, using methods described previously [58,59]. For the intact CYP116B1, the overlapping spectral contributions of the individual haem, FMN and 2Fe-2S cofactors resulted in difficulty in deconvoluting potentials for the three centres in the multicofactor enzyme. However, it is clear that the reduction process occurs in two phases. In the first phase (black spectra in Fig. 7A, completed in the range of 0 to −200 mV versus the normal hydrogen electrode [NHE]), spectral bleaching of the contributions of the iron–sulfur and FMN cofactors occurs at ∼ 400–600 nm (as seen for the dithionite-reduced spectrum in Fig. 2A). There is no obvious development of a long wavelength signal (at ∼ 600 nm) during this reductive phase (Fig. 7A), suggesting that there is negligible formation of the FMN neutral semiquinone, as also observed in the dithionite-reduced CYP116B1 EPR spectrum (Fig. 5). This may indicate that the reduction potential for the FMN semiquinone/hydroquinone couple (E2) is more positive than that for the oxidized/semiquinone couple (E1). With the assumption that absorption changes at 485 nm (in the potential range where there is negligible haem contribution) reflect primarily the transition from FMN quinone to hydroquinone (E12) (with some absorption change also resulting from the reduction of the 2Fe-2S cluster), the midpoint potential for the two-electron FMN reduction was estimated as −122 ± 9 mV (Fig. 7B). Data fitted at 409 nm (an isosbestic point for the haem iron Fe(III) to Fe(II) transition) produced a similar value (−134 ± 8 mV) for the combined reduction of the FMN/2Fe-2S centres. Although differing proportions of absorption contribution probably come from the FMN and iron–sulfur cluster at the two wavelengths, the midpoint potential values are within error and it is clear that complete reduction of both cofactors occurs in this potential range.
Figure 7. Spectroelectrochemical titration of CYP116B1. (A) UV-visible spectra recorded during a redox titration of CYP116B1 (6.3 μm). The arrows show directions of absorbance changes occurring at different wavelengths during the reductive phase of the titration. Absorbance spectra demonstrating the reduction of the FMN and 2Fe-2S clusters at more positive potentials are shown as black lines, while spectra showing mainly reduction of the CYP116B1 haem iron from ferric to ferrous (at more negative potentials) are shown as red lines. (B) A plot of absorbance data at 485 nm versus applied potential in the range from ∼ +25 to −200 mV (versus NHE). Data are fitted using the Nernst function to define a midpoint potential of −122 ± 9 mV for the reduction of both the FMN and 2Fe-2S centres in CYP116B1. (C) A plot of absorbance data at 418 nm versus applied potential in the range from ∼ +25 to −450 mV (versus NHE). The plot shows two absorbance transitions, and data were fitted using a two-electron Nernst function to define midpoint potentials of 160 ± 9 mV (encompassing both FMN and 2Fe-2S centres) and −301 ± 7 mV (for the CYP116B1 haem iron Fe(III)/Fe(II) couple).
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In the second stage of CYP116B1 reduction that occurs in the range from −200 to −500 mV versus NHE, the haem iron is converted from the ferric to the ferrous state. Haem iron reduction occurs with a shift in the Soret peak from 418 to 410 nm (red spectra in Fig. 7A), consistent with the retention of a cysteinate-ligated CYP116B1 haem iron and similar to the spectral maximum observed for the reduced form of P450 BM3 . Data fitted at 393 nm, where there are negligible absorbance changes from the early part of the reductive (FMN and iron-sulfur) titration, show a single transition with a midpoint potential of −291 ± 9 mV (data not shown). Absorption versus potential data plots at the oxidized Soret maximum (418 nm) show a clear two-stage transition (Fig. 7C). The decrease in A418 intensity in the first stage of the titration is attributed to the reduction of the FMN and iron-sulfur cofactors. This accounts for approximately half of the overall absorption change and is indicative of high levels of incorporation of these cofactors in CYP116B1. The midpoint potentials for the two transitions were determined using a two-electron Nernst function , yielding values of (a) −160 ± 9 mV to encompass the FMN E12 and iron–sulfur cluster [2Fe-2S]2+/[2Fe-2S]+ couples and (b) −301 ± 7 mV for the haem iron Fe(III)/Fe(II) couple. These midpoint potential values are consistent with the data fits at other wavelengths. Variations in the relative optical contributions of the FMN and 2Fe-2S cofactors may explain the small differences in redox-potential estimates for their combined reduction at different analysis wavelengths.
Identification of CYP116B1-catalysed reaction products of thiocarbamate herbicides
Following the fatty acid/thiocarbamate binding and steady-state kinetic studies reported above, we undertook experiments to establish whether CYP116B1 catalysed the oxidation of unsaturated fatty acids or of the thiocarbamates vernolate and EPTC. No formation of oxidized products was observed for NAD(P)H-driven turnover of CYP116B1 with monounsaturated fatty acids. However, preliminary studies indicated that products were formed from both vernolate and EPTC, with superior levels of product formation seen using NADPH as the reducing coenzyme (consistent with the steady-state kinetic data in Table 1). A detailed characterization of the products from the NADPH-dependent turnover of CYP116B1 with these thiocarbamates was thus undertaken by liquid chromatography (LC)-MS, as described in the Materials and methods.
HPLC was used to analyse products from CYP116B1-catalysed oxidation of vernolate and revealed major features in the chromatogram at 34–39 min after sample loading, in addition to the remaining parent molecule that eluted between 50 and 52 min (Fig. 8A). The new features consisted of two major product peaks, with a further small peak distinguishable between these major peaks. These product features are absent from vernolate-only or enzyme-only controls. The shorter retention time of these peaks indicates that they arise from compounds more polar in nature than the parent vernolate. The mass spectra of the major bands at 37.1 and 38.4 min in the chromatogram are nearly identical, with two major peaks observed with m/z values at 220 and 144 (Fig. 8B). The larger peak, at m/z 220, corresponds to the mass (M+H) of vernolate with a single hydroxylation. Further analysis of this peak performed by collision-induced disassociation (MS2) identified a single MS2 fragment with m/z ratio 144. Analysis of the vernolate structure revealed this MS2 fragment to be a hydroxylated form of vernolate lacking the S-propyl group and probably arising from cleavage of the S-C bond (Fig. 8C). MS2 analysis of the secondary HPLC-derived peak at m/z 144 revealed a single feature with an m/z ratio of 116, which again is a hydroxylated product of a vernolate fragment lacking both the S-propyl group and the carbonyl group (Fig. 8D). This fragmentation pattern that shows loss of the S-propyl group is consistent with that observed in the vernolate-only controls (Fig. S2A,B) and confirms the position(s) of CYP116B1-catalysed hydroxylation to be on one of the N-propyl chains. The fact that we observed two major peaks in the HPLC chromatogram with an almost identical mass spectrum may also be indicative of hydroxylation occurring at different positions on the N-propyl chain, although we cannot define their exact position(s) from these data. The mass spectrum of the third small species observed at 37.7 min in the chromatogram (indicated by an arrow in Fig. 8A) gave a distinct single peak with an m/z of 162 (Fig. 8E). This m/z ratio is consistent with dealkylation (loss of a propyl group) from the parent vernolate. This is most likely to be an N-dealkylation reaction given the different m/z ratio to that observed for the loss of the S-propyl group upon fragmentation. However, this dealkylated product is a minor species compared with the hydroxylated form(s) of vernolate, as seen from the difference in sizes of the respective peaks in the HPLC chromatogram. Similar hydroxylated products on the N-propyl chains are also observed for the thiocarbamate EPTC, which differs by a CH2 group in the S-ethyl chain (S-propyl in vernolate) (Fig. S3), although these are found in lower amounts compared with the vernolate products. There is no evidence of CYP116B1-mediated N-dealkylation with EPTC, although we cannot rule out small amounts of such a product. Given the greater proportion of hydroxylation and N-dealkylation seen with vernolate over EPTC, it appears likely that CYP116B1 binds more favourably to the S-propyl group in vernolate than to the S-ethyl group in EPTC to enable oxidative catalysis.
Figure 8. Analysis of products of CYP116B1-catalysed vernolate oxidation by liquid chromatography (LC)-MS. (A) HPLC chromatogram showing elution profiles of vernolate substrate (upper blue line) and CYP116B1-catalysed oxidation products of vernolate (lower black line). Vernolate elutes as a single peak between 50 and 52 min. Two additional peaks are observed following turnover with CYP116B1. These elute between 34 and 39 min and were determined to be hydroxylated forms of vernolate. A further sharp peak of lower intensity is also identified within this region, eluting at 37.7 min and indicated by the arrow. This feature is assigned to an N-dealkylated product of vernolate. (B) The mass spectrum of the chromatogram peaks at 37.1 and 38.4 min (both found to contain almost identical masses) showing two major species with m/z values of 220 and 144, and a smaller m/z feature at 162. The m/z 220 species results from hydroxylation of vernolate (m/z = 204 + 16). The structure of a potential hydroxylated product of vernolate is shown as an inset. (C) MS2 analysis of the m/z 220 species with a resultant m/z at 144. This 144 m/z species is assigned to a hydroxylated fragment of vernolate, with the S-propyl group removed. A representative structure of the hydroxylated vernolate fragment structure is shown as an inset. (D) MS2 analysis of the m/z 144 species with a resultant m/z at 116. This is assigned to a hydroxylated fragment of vernolate, with the S-propyl and adjacent carbonyl groups removed. These data are further confirmatory that the hydroxylation occurs on one of the N-propyl chains, but do not define an exact position. A representative structure of this hydroxylated fragment is shown as an inset. (E) Mass spectrum of the minor peak at 37.7 min in the ‘CYP116B1 + vernolate’ chromatogram (A), showing a predominant m/z peak at 162. This is consistent with dealkylation (loss of a propyl group) from vernolate. While the dealkylation could theoretically occur with removal of either an N- (shown inset) or an S-propyl chain, the likely product is the N-dealkylated form (as seen for the reaction of Rhodococcus erythropolis ThcB with EPTC), occurring by CYP116B1-catalysed C-N bond cleavage following successive hydroxylations at adjacent carbons on the N-propyl chain .
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