Structural insights into S-lignin O-demethylation via a rare class of heme peroxygenase enzymes

Harlington, Alix C.; Das, Tuhin; Shearwin, Keith E.; Bell, Stephen G.; Whelan, Fiona

doi:10.1038/s41467-025-57129-6

Download PDF

Article
Open access
Published: 20 February 2025

Structural insights into S-lignin O-demethylation via a rare class of heme peroxygenase enzymes

Nature Communications volume 16, Article number: 1815 (2025) Cite this article

7044 Accesses
5 Citations
104 Altmetric
Metrics details

Subjects

Abstract

The O-demethylation of lignin aromatics is a rate-limiting step in their bioconversion to higher-value compounds. A recently discovered cytochrome P450, SyoA, demethylates the S-lignin aromatic syringol. In this work, we solve high-resolution X-ray crystal structures of substrate-free and substrate-bound SyoA and evaluate demethylation of para-substituted S-lignin aromatics via monooxygenase and peroxide shunt pathways. We find that SyoA demethylates S-lignin aromatics exclusively using the peroxide shunt pathway. The atomic-resolution structures reveal the position of non-canonical residues in the I-helix (Gln252, Glu253). Mutagenesis of this amide-acid pair in SyoA shows they are critical for catalytic activity. This work expands the enzymatic toolkit for improving the capacity to funnel lignin derived aromatics towards higher value compounds and defines the chemistry within the active site of the enzyme that enables peroxygenase activity. These insights provide a framework for engineering peroxygenase activity in other heme enzymes to generate easier to use biocatalysts.

Highly stable and tunable peptoid/hemin enzymatic mimetics with natural peroxidase-like activities

Article Open access 31 May 2022

Resolving the metabolism of monolignols and other lignin-related aromatic compounds in Xanthomonas citri

Article Open access 12 September 2024

Critical enzyme reactions in aromatic catabolism for microbial lignin conversion

Article 25 February 2022

Introduction

Ubiquitous in all domains of life, the superfamily of heme-dependent cytochrome P450 enzymes (P450s) catalyze selective oxidations of unactivated C−H bonds. In recent years, P450s have gained traction as catalysts for lignin valorization—the process of converting the plant polymer lignin into commercially valuable chemicals¹. The CYP255 family of P450s; so far consisting of GcoA², AgcA³, and SyoA⁴, catalyze the O-demethylation of lignin monoaromatics, which is an important step in the valorization of lignin. Lignin is a complex heterogeneous polymer comprising p-coumaryl (H-type), coniferyl (G-type) and sinapyl (S-type) alcohol subunits, which have zero, one or two methoxy groups, respectively⁵ (Fig. 1a). Aromatics derived from lignin depolymerization must be O-demethylated to diols to permit formation of ring-opened compounds, which can be funnelled to high-value compounds, e.g., organic acids (cis,cis-muconate, pyruvate and lactate); pyridine-related organic acids (pyridine-2,4-dicarboxylic acid); and lipids which can be used to make various materials (polyamides and polyesters), fuels and flavors⁶. So far, three classes of enzymes have been identified that catalyze O-demethylation of S-lignin aromatics including: tetrahydrofolate (THF)-dependent demethylases^7,8, Rieske non-heme iron-dependent oxygenases^9,10, and P450s⁴ (Fig. 1b). P450 enzymes have been engineered for improved demethylation of syringol, including GcoA¹¹ and CYP102A1 (P450_BM3)¹²; however, native enzymes capable of the O-demethylation of S-lignin aromatics remain largely undescribed.

**Fig. 1: Lignin structure, S-lignin demethylases and P450 catalytic cycle.**

Most P450s function as monooxygenases requiring redox partners to transfer two electrons to the heme iron from the nicotinamide cofactor NAD(P)H to activate dioxygen (O₂)^13,14. Activation of dioxygen results in the formation of the oxy-ferryl porphyrin radical cation species compound I (Cpd I), the reactive intermediate^15,16 (Fig. 1c). In many P450s this process is mediated by two residues within the oxygen binding groove of the I-helix—referred to as the acid-alcohol pair (Asp/Glu-Thr/Ser). Specifically, the acid aids in the delivery of protons to the heme-oxygen intermediates, while the alcohol is believed to stabilize the ferric-hydroperoxo intermediate, compound 0 (Cpd 0) for efficient formation of Cpd I^{17,18,19,20,21}. P450s can also use H₂O₂ to generate Cpd I by using the “peroxide shunt pathway”, circumventing the need for additional redox partner proteins and expensive NAD(P)H cofactor (Fig. 1c)^22,23. The majority of P450s are unable to use H₂O₂ efficiently, leading to heme destruction and enzyme inactivation²⁴. However, some families of heme enzymes possess naturally high levels of peroxygenase activity and deploy residues capable of acid-base chemistry to activate H₂O₂. For example, the CYP152 family use the substrate’s carboxyl group, coordinated to an Arg, to function as peroxygenases^25,26,27. Furthermore, the P450 (SscaCYP) was shown to hydroxylate trans-β-methyl-styrene and catalyze the sulfoxidation of thioanisole using H₂O₂²⁸. Structural analysis of SscaCYP revealed two acid-base residues in the active site; an Asp which replaces the canonical I-helix Thr, and a Glu near the K-helix. Other heme-containing peroxidases and peroxygenases also deploy residues capable of acid-base chemistry on the distal side of the heme^22,29. It is believed that binding of H₂O₂ to the ferric form of these enzymes results in deprotonation of the iron-bound oxygen, forming Cpd 0, with a protonation step resulting in O–O bond scission and dehydration leading to Cpd I (Fig. 1c).

Here we report high-resolution X-ray crystal structures of the recently identified CYP255 enzyme SyoA from Amycolatopsis thermoflava N1165 in the open substrate-free and closed substrate-bound states. These structures collectively reveal capacity in the active site to accommodate O-demethylation of para-substituted S-lignin aromatics. We demonstrate that SyoA functions more efficiently using the peroxide shunt pathway to demethylate S-lignin aromatics compared to the monooxygenase pathway. Additionally, we investigate the role of the amide-acid pair (QE) of residues, which sit above the heme within the I-helix, in the peroxygenase activity of the CYP255 enzymes SyoA and GcoA and monooxygenase activity of GcoA. Mutagenesis of the QE residues in SyoA and GcoA illustrates that, for this residue pair, both monooxygenase and peroxygenase activities are profoundly impacted by reversion to the canonical P450 ET sequence.

Results

Structural analysis and conformational change of SyoA

To investigate syringol binding by SyoA, we performed structural characterization by X-ray crystallography. Crystal structures of substrate-free (PDB: 8u09) and syringol-bound (PDB: 8u19) SyoA were determined at resolutions of 1.98 and 1.26 Å, respectively (Fig. 2, Supplementary Figs. 1, 2a–b, Supplementary Tables 1–4).

**Fig. 2: X-ray crystal structures of SyoA in the open substrate-free and closed substrate-bound states.**

SyoA exhibits the characteristic single-domain trigonal prism shape, consisting of sixteen α-helices and four β-sheets, with the central heme iron coordinated by Cys360 (Fig. 2a, d, Supplementary Fig. 1). The substrate-free enzyme adopts an open conformation, with a water-filled channel that exposes the active site and heme to solvent (Fig. 2a–c, Supplementary Fig. 2a). The substrate access channel consists of a network of 13 hydrogen-bonded waters (Supplementary Fig. 3). On the distal side of the heme, an axial water is coordinated ~2.5 Å from the heme iron (Fig. 2b). Furthermore, unidentified electron density was observed in the active site that was too large for water and too small for syringol. The best candidate for the densities above the heme and Phe399 was glycerol from the cryoprotectant, while density near His75 was consistent with acetate present in the crystallization conditions (Supplementary Fig. 3).

In contrast, the syringol-bound structure of SyoA adopts a closed conformation with the heme and substrate buried within a closed active site (Fig. 2d, Supplementary Fig. 2b). The active site of SyoA is predominantly hydrophobic, consisting of His75, Leu79, Leu85, Ile173, Val245, Val296, Ala299, Ser300, and Phe399 (Fig. 2e, f). Syringol is situated in the active site, with one methoxy carbon positioned towards the heme ~3.6 Å from the iron, and the other methoxy group positioned adjacent to His75 and Ile173 (Fig. 2e). Moreover, the aromatic ring of syringol is held between Leu85 and Phe399 with the hydroxyl group of syringol hydrogen-bonded to the carbonyl of Val245 (Fig. 2e–f). The network of water molecules observed in the substrate-free structure are not present in the substrate-bound form.

The conformational change between the open and closed state of SyoA is characterized by an ~19° displacement of helices B′, F, G, and H; and the F/G and H/I loops, via a hinge-like mechanism (Fig. 2g, Supplementary Fig. 4a and Supplementary Table 2). Displacements are observed for residues His75 (2.5 Å) and Leu79 (2.6 Å) of the B′-helix and Ile173 (4.4 Å) of the F-helix, which directly interact with one of the methoxy moieties of syringol (Fig. 2h, Supplementary Table 3). The G/H-loop, comprising residues 206–221, was not modeled in the open conformation due to poor resolution of the electron density. The average B-factors were highest surrounding the G/H-loop and were significantly lower in the closed state, consistent with stabilization of this region upon substrate binding (Supplementary Fig. 4b). Clear electron density was observed in the closed conformation enabling building and refinement of the G/H-loop in the bound state.

The closest structural homolog to SyoA is GcoA (PDB: 5ncb)²; the aligned structures reveal that relative to GcoA, there is additional space in the active site of SyoA, consistent with SyoA being permissive of binding to S-lignin aromatics which contain an additional methoxy group compared to G-lignin aromatics (Supplementary Figs. 5, 6). A triad of phenylalanine residues (Phe75, Phe169, and Phe395) is critical in positioning the aromatic ring of guaiacol in the active site of GcoA². SyoA does not contain this triad of phenylalanine residues; instead, Leu79, Ile173 and Phe399 occupy equivalent positions in the active site (Supplementary Fig. 6a, b). Unique to the active site of SyoA is His75 (the equivalent residue in GcoA is Gly71), which partially fills the void in the active site occupied by larger aromatic residues in GcoA (Supplementary Fig. 6a).

Active site binding of para-substituted S-lignin aromatics

Substrate binding to P450s commonly induces a shift from a low to high-spin (HS) ferric state. The low-spin (LS) state is characterized by heme coordination of 6 ligands (Soret maximum at ~419 nm), including the active site water; the transition to a HS state (Soret maximum at ~390 nm) occurs when the substrate displaces this water. Previously, our analysis showed that SyoA preferentially binds syringol (1) over guaiacol⁴. The high-resolution crystal structure of syringol-bound SyoA reveals additional space in the active site at the substrate C4 carbon – para to the hydroxyl group of syringol (Fig. 3a, b). Consequently, we investigated binding of para-substituted S-lignin aromatics to SyoA including 4-methylsyringol (2), syringaldehyde (3), syringic acid (4), acetosyringone (5), 4-allylsyringol (6), sinapic acid (7) and sinapyl alcohol (8) (Fig. 4a). Strong absorbance of syringaldehyde, acetosyringone and sinapic acid in the visible spectrum from 420–390 nm made it difficult to investigate binding using the Soret band: therefore, the Q-bands at 568 and 535 nm were used to interrogate binding. Among the tested S-type aromatics, four demonstrated binding to SyoA with syringol and 4-methylsyringol inducing a ~70% shift to the HS form and 4-allylsyringol a ~15% shift using the Soret band (Fig. 4a). Addition of syringaldehyde to SyoA caused a decrease in the Q-bands, and an increase in the charge transfer band at ~640 nm indicative of ligand binding in the active site (Fig. 4a and Supplementary Fig. 7a). Binding analysis showed that SyoA binds 4-methylsyringol with the highest affinity (K_D = 7.2 ± 0.2 μM), followed by syringol (K_D = 16 ± 1 μM), 4-allylsyringol (K_D = 151 ± 11 μM) and syringaldehyde (K_D = 341 ± 25 μM) (Fig. 4b–e and Supplementary Fig. 7a–d).

**Fig. 3: Comparison of active site volumes in the closed substrate-bound state.**

**Fig. 4: Binding analysis of S-lignin aromatics to SyoA.**

Crystallization of SyoA was subsequently conducted with the addition of 4-methylsyringol. The atomic-resolution crystal structure of SyoA bound to 4-methylsyringol (PDB: 8u1i) was solved at 1.12 Å; no significant differences were observed in the active site, nor in the position of the substrate, with an all-atom RMSD of 0.35 Å between the syringol and 4-methylsyringol-bound structures (Fig. 3c, d, Supplementary Figs. 2c, 8 and Supplementary Table 1). The 4-methyl group occupies the vacant space observed in the syringol-bound structure, with the remaining space occupied by three water molecules which are conserved in both structures (Fig. 3c, d).

SyoA O-demethylates S-lignin aromatics using hydrogen peroxide

The CYP255 members GcoA and AgcA catalyze the O-demethylation of G-lignin aromatics, as a two-component system, using their respective three-domain redox partners (GcoB and AgcB). To determine the catalytic range of SyoA, we first investigated the O-demethylation of S-lignin aromatics using the monooxygenase pathway with the predicted redox partner SyoB, the encoding gene being adjacent SyoA (Supplementary Fig. 9). In contrast to GcoB and AgcB, SyoB consists of a predicted N-terminal FMN binding domain, NADH binding domain and a 2Fe-2S ferredoxin domain making it more structurally similar to the phthalate dioxygenase reductase family³⁰. Unlike the other CYP255 members, the reconstituted SyoA/SyoB monooxygenase system did not O-demethylate syringol, 4-methylsyringol, or syringaldehyde (Supplementary Figs. 10, 11). However, NADH was consumed in the reaction containing SyoA, SyoB and syringol (Supplementary Fig. 10a–c). HPLC analysis identified 2,6-dimethoxyhydroxyquinone (1a) as the major product which, importantly, was also observed in the negative control excluding SyoA. This result indicates that syringol is reduced by SyoB in the presence of NADH, leading to uncoupling of electron transfer from SyoA (Supplementary Fig. 10d).

We next investigated the O-demethylation of syringol, 4-methylsyringol, syringaldehyde and 4-allylsyringol using the peroxide shunt pathway. Reactions with syringol, 4-methylsyringol and syringaldehyde all yielded demethylated products (catechols) and formaldehyde (9) as a by-product (Fig. 5, Supplementary Fig. 12). We observed no evidence that SyoA could do two successive demethylation reactions to produce pyrogallols. Additionally, several other products (due to overoxidation of the catechols by excess hydrogen peroxide) were observed in each reaction. The O-demethylation of syringol (1) yielded 142 ± 15 μM (total turnover number (TTN) 142) 3-methoxycatechol (1b), with ~65% of the substrate consumed over 60 min (Fig. 5, Supplementary Figs. 13, 17a). The demethylation of 4-methylsyringol (2) occurred to the greatest degree, with 795 ± 30 μM (total turnover number (TTN) 795) 3-methoxy-5-methylbenzene-1,2-diol (2a) produced over the 60 min reaction (Fig. 5, Supplementary Figs. 14, 17b). Syringaldehyde was demethylated to 5-hydroxyvanillin (3a) (308 ± 27 μM, TTN 308) (Fig. 5, Supplementary Fig. 15a). The reaction with syringaldehyde also liberated 2,6-dimethoxyhydroxyquinone (1a) / 2,6-dimethoxy-1,4-benzoquinone (3c) and 1,3,4-trihydroxy-5-methoxybenzene (3b) due to oxidation of the aldehydes by hydrogen peroxide via a Dakin oxidation reaction³¹ (Supplementary Figs. 15b, c, 17c, 18). The oxidation of 4-allylsyringol (6) by SyoA yielded several metabolites in low yield as observed by HPLC, with one of the products confirmed to be 4-allylsyringol +[O] (m/z = 210) (Supplementary Figs. 16, 17d).

**Fig. 5: O-demethylation of S-lignin aromatics by SyoA using the peroxide shunt.**

Structure and mutagenesis of the I-helix catalytic residues

The I-helix of P450 enzymes contains residues important for oxygen activation and catalysis³². We previously identified that the I-helix of the CYP255 family does not have the conserved acid-alcohol pair believed to be responsible for the monooxygenase function of most P450 enzymes⁴. The crystal structures of SyoA and GcoA reveal that the glutamine-glutamate pair (Gln252 and Glu253 SyoA; Gln248 and Glu249 GcoA) are observed in a structurally equivalent position to the acid-alcohol residues of most P450 enzymes and are likely to be important for the peroxygenase activity of the CYP255 family (Fig. 6a–c).

**Fig. 6: Mutagenesis of the I-helix catalytic residues.**

In the substrate-free state of SyoA, the I-helix exhibits a break in the α-helical structure from Gly249 to Gln252 due to the backbone amide of Glu253 forming a hydrogen bond with the carbonyl oxygen of Gly250 (Fig. 6a). In the syringol-bound complex, the break is resolved due to the backbone amide of Glu253 forming a hydrogen-bond with the carbonyl of Gly249 instead (Fig. 6b). This conformational change affects the position of the glutamine–glutamate pair. In the substrate-free structure Gln252 is hydrogen-bonded to Glu141 of the E-helix, while Glu253 forms several interactions with surrounding solvent molecules and His256 (Fig. 6a). Upon substrate binding, Gln252 forms a hydrogen bond to Glu253, moving closer to the heme (Fig. 6b). Additionally, the direct hydrogen bond between Glu253 and His256 is disrupted by a water molecule. The I-helix of GcoA (PDB: 5ncb) adopts a similar conformation to SyoA, however, Gln248 hydrogen bonds to Asn188 of the F-helix and Glu249 directly interacts with His252 (Fig. 6c). In the substrate-bound structures the glutamic acid is located ~7.4 Å (SyoA) or ~8.1 Å (GcoA) from the heme iron and ~6.0 Å (SyoA) or ~6.8 Å (GcoA) from the 2-methoxy carbon.

To determine whether the non-canonical I-helix residues of the CYP255 family are important for peroxygenase activity, we performed site-directed mutagenesis on SyoA (Gln252 and Glu253) and GcoA (Gln248 and Glu249). We investigated double mutants (SyoA_ET and GcoA_ET), where the QE residues were replaced with those commonly found in typical P450 monooxygenases. Additionally, to assess the specific contribution of each individual residue to catalytic activity, we also created single mutants (SyoA_EE/GcoA_EE and SyoA_QT/GcoA_QT). UV-Vis analysis demonstrated that binding of syringol to SyoA and guaiacol to GcoA was not drastically impacted by the mutations, indicating that changes in catalytic activity are not likely due to loss of substrate binding (Supplementary Fig. 19a–b). Mutation of either the glutamine or glutamate in SyoA and GcoA was sufficient to completely remove catalytic function using the peroxide shunt (Fig. 6d–e). Furthermore, we also investigated the impact of the mutations on the monooxygenase activity of GcoA, which showed a significant reduction in the monooxygenase activity for GcoA_EE and GcoA_ET and complete loss in activity for the GcoA_QT mutant (Supplementary Fig. 20).

Discussion

In contrast to chemical catalytic approaches, the O-demethylation of lignin aromatics using biocatalysis is of great interest as it offers an environmentally friendly method for converting lignin to high-value compounds^33,34. Few S-lignin specific O-demethylases have been characterized, limiting the bioconversion of lignin from hardwoods and grasses. In this study, we determined the X-ray crystal structures of CYP255 SyoA in the open substrate-free and closed substrate-bound states and show that, in vitro, SyoA only catalyzes O-demethylation using the peroxide shunt pathway.

Compared to GcoA, SyoA has additional space in the active site allowing for binding of larger S-lignin aromatics. Specifically, SyoA contains an isoleucine (Ile173) at the equivalent position of a key phenylalanine residue (Phe169) present on the F-helix, which has been shown to prevent productive binding of syringol in GcoA due to a steric clash with the second methoxy group¹¹. Mutagenesis of this residue in GcoA to an alanine increases specificity for syringol¹¹ demonstrating that Ile173 in SyoA and Phe169 in GcoA may be responsible for determining specificity between S-lignin and G-lignin aromatics, respectively. Compared to GcoA, the active site of SyoA contains additional space at the para position allowing for binding of para-substituted aromatics (4-methylsyringol, syringaldehyde and 4-allylsyringol), due to the presence of an active site serine (equivalent position in GcoA features a threonine). Syringaldehyde is a major product of oxidative lignin depolymerization^35,36, while syringol and 4-methylsyringol are more commonly observed during lignin pyrolysis^36,37, making SyoA a suitable catalyst for aromatics derived from different methods of lignin depolymerization.

Various P450 enzymes are known to undergo conformational changes from open to closed states upon substrate binding³⁸. In this work, we provide structural evidence that SyoA switches from an open to closed conformation in the presence of substrate, with relocation of the F, G and B′ helices to establish the closed hydrophobic active site of SyoA. The results described here support molecular dynamics (MD) simulations that show closing of the active site of GcoA in response to substrate binding². Contrary to these observations, MD simulations predict AgcA (CYP255A1) prefers a closed conformation with smaller substrates but adopts an open conformation with larger substrates³⁹.

Like most P450 monooxygenase enzymes, GcoA and AgcA catalyze substrate oxidation using their respective redox partners, NAD(P)H and O₂. Interestingly, SyoA with its predicted redox partner SyoB demonstrated no O-demethylation of syringol, 4-methylsyringol and syringaldehyde. However, low levels of SyoB-dependent oxidation of syringol to 2,6-dimethoxyhydroxyquinone was observed. The presence of redox-active quinones can result in NADH oxidation, transferring the electrons via the redox partners to the quinone. This results in uncoupling of electron transfer from the P450 activity, explaining the oxidation of NADH without product formation⁴⁰. This phenomenon has also been observed for VanB and syringic acid¹⁰ indicating that some lignin aromatics may lead to uncoupling via redox cycling. However, SyoA and GcoA can efficiently use the peroxide shunt pathway to catalyze O-demethylation of their substrates—an uncommon feature of P450s⁴. Using the peroxygenase activity of SyoA, we were able to show O-demethylation of syringol, 4-methylsyringol and syringaldehyde. To our knowledge this is the first report of an enzyme that can demethylate the S-lignin aromatics 4-methylsyringol and syringaldehyde—which present as key bottlenecks to biological lignin valorization.

SyoA stands out among lignin-degrading enzymes for its ability to catalyze the O-demethylation of S-lignin aromatics using H₂O₂, a feature more common with certain heme-containing peroxidases than with other enzyme classes. Laccases, for instance, utilize O₂ to perform indirect C-C bond cleavage in lignins by generating mediators, such as phenoxy radicals, which facilitate further degradation^6,41. In contrast, heme-containing peroxidases, including manganese peroxidases, lignin peroxidases, versatile peroxidases, and dye-decolorizing peroxidases, employ H₂O₂ to directly cleave C-C bonds in lignin structures⁴¹. Compared to hydroxylases, decarboxylases, and dioxygenases—which commonly use O₂ for their catalytic activity—the CYP255 family, including SyoA, is distinct in its efficient use of H₂O₂, thereby simplifying the catalytic requirements⁴².

High-resolution structural analysis of the I-helix of SyoA has revealed the conserved acid-alcohol pair—typically found in P450 monooxygenases—is replaced with an amine-acid pair (glutamine and glutamate). Interestingly, peroxidases and peroxygenases contain amino acids capable of acid-base chemistry (histidine, aspartate, or glutamate) on the distal side of the heme^22,29. We performed site-directed mutagenesis on the amine-acid pair of SyoA and GcoA and demonstrated the importance of the non-canonical I-helix residues for both the monooxygenase and peroxygenase function. All mutants lost the ability to use the peroxide shunt pathway with significant reduction in the monooxygenase activity also observed for GcoA, demonstrating the amide-acid pair is important for both oxygen and hydrogen peroxide activation in the CYP255 family.

We postulate that the CYP255 family may have evolved hybrid monooxygenase/peroxygenase enzyme activity for several reasons. Phylogenetic analysis of the CYP255 family compared to other P450 families shows that the peroxygenase activity may have evolved from a common ancestor to the CYP255 and CYP1232 family – both of which demethylate aromatics (Supplementary Fig. 21). Additionally, the peroxygenase activity appears to have evolved separately compared to other P450 peroxygenases including the CYP152 and SscaCYP (CYP177), indicating unique selection pressures. As shown by the reaction of syringol with SyoB, CYP255 enzymes may have evolved peroxygenase activity to avoid uncoupling via redox cycling by quinones. Hydroquinones including 2-methoxyhydroquinone (MBQH₂) and 2,6-dimethoxyhydroxyquinone (DBQH₂) can be derived from products of lignin degradation^43,44. Furthermore, exposing lignin to light facilitates production of hydrogen peroxide^45,46. Interestingly, many enzymes involved in the depolymerization of lignin also use H₂O₂ as a cosubstrate, suggesting these peroxygenases may have evolved to use hydrogen peroxide as a ‘free’ catalytic driver, removing the reactive oxygen species and thereby protecting the P450 from oxidative stress⁴⁷. Thus, the CYP255 family may have evolved hybrid monooxygenase/peroxygenase enzyme activity as a mechanism to minimize uncoupling in the presence of quinones, using the hydrogen peroxide generated to drive catalysis.

The biochemical and structural data presented here suggest that SyoA is involved in the O-demethylation of S-lignin aromatics, which could allow Amycolatopsis thermoflava to funnel lignin monoaromatics to central carbon metabolism. Demethylation of syringol, 4-methylsyringol and syringaldehyde liberate catechols which other enzymes could further catabolize as an energy source to facilitate efficient lignin breakdown. Moreover, the atomic resolution structural insights also offer scope for engineering the active site of SyoA to accommodate other S-lignin aromatic substrates, including syringic acid or 3-O-methylgallic acid, expanding its utility. Additionally, the peroxygenase activity of the CYP255 family offers an inexpensive and clean approach to demethylation compared to traditional monooxygenases, which require specific redox partners and expensive nicotinamide cofactors. Using the sequence of the I-helix and surrounding regions of the CYP255 family may enable the retrofitting of peroxygenase activity into other P450s, making biocatalysis more favorable for larger scale applications. Indeed, the weak peroxygenase activity of CYP119⁴⁸ and CYP199, CYP154C8, and CYP102⁴⁹ was improved by mutating the acid-alcohol pair and surrounding residues to resemble the I-helix of the CYP255 enzymes.

Methods

General

Chemical reagents were purchased from Sigma-Aldrich. HPLC grade solvents were from Sigma-Aldrich, Ajax Finechem, and Chem-Supply. Enzymes, biochemical and molecular biology reagents were supplied by Sigma-Aldrich, New England Biolabs, and Thermo Fisher Scientific. Crystallography reagents, MicroMounts, MicroLoops, and MicroTools were from Hampton Research and Molecular Dimensions and the NVH oil from (Cargille Laboratories). Substrate and product stocks were prepared in DMSO. IS (internal standard) stock solutions were prepared in EtOH.

Gene cloning

A pET3a vector containing a codon optimized gene encoding SyoA was obtained using Gibson isothermal assembly⁵⁰, by 3-fragment assembly with a single polymerase chain reaction (PCR)-generated insert fragment containing the syoA gene and two plasmid backbone fragments. The syoA gene was PCR amplified from the pET29-SyoA vector⁴ using primers 3558 and 3559 (Supplementary Tables 5 and 6). The pET3a backbone fragments were PCR-generated using primers pairs 2281 and 3312 and 3497 and 1330 (Supplementary Table 6). SyoA mutants were made by 2-fragment Gibson assembly using the pET3a SyoA vector as the template. Primers 679 and 3991 were used to generate the first fragment and primer 1330 paired with 3992, 3993, or 3994 were used to generate the mutants SyoA_QT, SyoA_EE, SyoA_ET, respectively. Primers 215 and 315 (T7 promoter and T7 terminator, respectively) were used to sequence the insert fragment (Australian Genome Research Facility, Adelaide, Australia). A pET26-GcoA vector⁴ was used for bacterial expression of GcoA. GcoA mutants (GcoA_EE, GcoA_QT, GcoA_ET) were purchased (Twist Bioscience) cloned in the pET28a plasmid with an N-terminal 6x His tag and Tobacco etch virus (TEV) cleavage site followed by a NdeI restriction site and the sequence for the gene of interest. Codon optimized genes encoding SyoB (based on the NCBI reference sequence: WP_027930991) and GcoB were purchased as gBlocks from IDT. An NdeI restriction site was incorporated at the 5’ end, followed by a double stop codon and KpnI and HindIII restriction sites. PCR was performed on the gBlock DNA using primers RS0114865 5’ and RS0114865 His 3’ for SyoB and RS0109265 5’ and RS0109265 His 3’ for GcoB to add a 6x His tag to the C-terminus of the gene and inserted into a pET29b vector using the NdeI and HindIII restriction sites. The final sequence was confirmed by DNA sequencing (Australian Genome Research Facility, Adelaide, Australia).

Protein expression and purification of SyoA and GcoA

The pET3a vectors containing syoA, syoA_EE, syoA_QT or syoA_ET; the pET26 vector containing the gcoA gene; and the pET28a vectors containing gcoA_EE, gcoA_QT or gcoA_ET were transformed into Escherichia coli BL21 (DE3) competent cells. The cells were plated on lysogeny broth (LB) agar containing ampicillin (100 µg/ml) for the pET3a vectors and kanamycin (50 µg/ml) for the pET26 and pET28a vectors and incubated at 37 °C overnight (~16–20 h). Single colonies were used to inoculate a 50 mL LB starter culture containing the appropriate antibiotic and grown overnight (~16–20 h) at 37 °C with shaking (180 rpm). 2.5 L flasks containing 1 L LB with antibiotics were inoculated with 1:100 v/v starter culture and incubated, 37 °C with shaking (150 rpm). At an OD₆₀₀ of 2, 1% v/v ethanol and 0.02% v/v benzyl alcohol were added to induce chaperone expression⁵¹, and the cultures incubated at 18 °C with shaking 100 rpm. Additionally, 3 mL/L trace elements (Per liter: Na₂EDTA (20.1 g), FeCl₃.6H₂O (16.7 g), CaCl₂.H₂O (0.74 g), CoCl₂.6H₂O (0.25 g), ZnSO₄.7H₂O (0.18 g), MnSO₄.4H₂O (0.132 g), CuSO₄.5H₂O (0.10 g)) were added for cofactor incorporation. After 30 min, protein expression was induced with addition of IPTG (isopropyl β-D-1-thiogalactopyranoside) to a final concentration of 0.1 mM, and the culture incubated overnight ~20 h at 18 °C with shaking at 100 rpm. Cells were harvested by centrifugation (6200 × g, 15 min, 4 °C).

The cell pellet was resuspended in ice-cold lysis buffer (50 mM Tris, pH 7.5, 50 mM NaCl, 20 mM imidazole). Cells were lysed by sonication (30 cycles of 15 s on, 45 s off, 70% amplitude, 19 mm probe, Sonics Vibra-Cell) and cell lysate clarified by centrifugation (40,000 × g, 20 min, 4 °C). Proteins were purified by immobilized nickel affinity chromatography (HisTrap HP, 5 mL column Cytiva) using a linear gradient of 20–250 mM imidazole at a flow rate of 5 mL/min. The samples were buffer exchanged into 50 mM Tris, pH 7.5 by centrifugal ultrafiltration using a Vivaspin 20 centrifugal concentrator (Sartorius) with a 10 kDa molecular weight cut-off. Wild type SyoA and GcoA were further purified by anion exchange (HiTrap Q FF, 5 mL Cytiva) using a linear gradient of 0–500 mM NaCl at a flow rate of 5 mL/min. Purified protein was concentrated by ultrafiltration to <10 mL and stored in 40–50% v/v glycerol at −20 °C. Glycerol was removed before use using a PD-10 desalting column (Cytiva) equilibrated in 50 mM Tris, pH 7.5. The concentration of each protein was estimated using the following extinction coefficients: SyoA (ε_417nm = 123 mM⁻¹ cm⁻¹)⁴, GcoA (ε_417nm = 119 mM⁻¹ cm⁻¹)⁴.

Protein expression and purification of SyoB and GcoB

The pET29b vectors containing syoB or gcoB were transformed into Escherichia coli BL21 (DE3) competent cells. The cells were plated on LB agar containing kanamycin (50 µg/ml) and incubated at 37 °C overnight (~16–20 h). Cells were grown as described above in 2.5 L flasks containing 1 L LB with antibiotics. At an OD₆₀₀ of 0.6, 1% v/v ethanol and 0.02% v/v benzyl alcohol were added and the cultures incubated at 18 °C with shaking 80 rpm. Additionally, 1 mM L-cysteine and 0.5 mM ferric ammonium citrate were added to aid cofactor incorporation. After 30 min, protein expression was induced with addition of 0.1 mM IPTG, and the culture incubated overnight ~20 h at 20 °C with shaking at 80 rpm. Cells were harvested by centrifugation (6200 × g, 15 min, 4 °C).

The cell pellet was resuspended in ice-cold lysis buffer (50 mM Tris, pH 7.5, 500 mM NaCl, 20 mM imidazole, 1 mM DTT) for SyoB and (50 mM Tris, pH 7.5, 50 mM NaCl, 1 mM DTT) for GcoB. Cells were lysed by sonication (30 cycles of 15 s on, 45 s off, 70% amplitude, 19 mm probe, Sonics Vibra-Cell) and cell lysate clarified by centrifugation (40,000 × g, 20 min, 4 °C). SyoB was purified by immobilized nickel affinity chromatography (HisTrap HP, 5 mL column Cytiva) using a linear gradient of 20–250 mM imidazole at a flow rate of 5 mL/min. GcoB was purified by ion-exchange chromatography, using a DEAE Sepharose column and characterized by UV-Vis absorbance spectroscopy (Supplementary Fig. 9). A linear gradient of 50–500 mM NaCl was used to elute the protein at a flow rate of 5 mL/min. The samples were buffer exchanged into 50 mM Tris, pH 7.5 by centrifugal ultrafiltration using a Vivaspin 20 centrifugal concentrator (Sartorius) with a 10 kDa molecular weight cut-off. Purified protein was concentrated by ultrafiltration to <10 mL and stored in 40–50% v/v glycerol at -20 °C. Glycerol was removed before use using a PD-10 desalting column (Cytiva) equilibrated in 50 mM Tris, pH 7.5. The concentration of SyoB and GcoB were estimated using the extinction coefficient (ε_423nm = 25.2 mM⁻¹ cm⁻¹)².

Protein crystallization and structure determination

For crystallization, the N-terminal His₆ tag was cleaved using His₆-TEV protease and separated from His₆-tagged SyoA, His6-tag and TEV protease by IMAC. SyoA was further purified by size exclusion chromatography (HiLoad 16/600 Superdex 200 pg, Cytiva) and concentrated to ~15 mg mL⁻¹ in 20 mM Tris pH 7.5 by ultrafiltration. Crystals were grown by sitting drop vapor diffusion at 289 K using 1 µL of protein with 1 µL of reservoir solution, including PEG/Ion I&II, Index, and PGA screens. SyoA in the substrate-free state crystallized within 1 week in conditions comprising 0.25 mM syringol (from a 50 mM stock in DMSO), 0.1 M sodium acetate pH 5, 5% w/v γ-PGA (γ-polyglutamic acid) and 12% w/v PEG 8000. Crystals were cryoprotected in reservoir solution supplemented with 25% v/v glycerol prior to flash cooling in liquid nitrogen. SyoA in the substrate-bound state crystallized within 1 week in conditions comprising 20 mM substrate (from a 200 mM stock in 40% DMSO), 0.2 M magnesium nitrate hexahydrate pH 5.9, 20% w/v PEG 3350 or 0.2 M lithium nitrate pH 7.1, 20% w/v PEG 3350. Crystals were cryoprotected in NVH oil prior to flash cooling in liquid nitrogen.

Diffraction data were collected for the substrate-free and syringol-bound crystals of SyoA at the MX1 beamline at the Australian Synchrotron⁵². Diffraction data were collected remotely for the 4-methylsyringol-bound crystal of SyoA at the MX2 beamline at the Australian Synchrotron using the Blu-Ice GUI interface to EPICS control and qeGUI⁵³. 360° of data were collected using 0.1° oscillations at a wavelength of 0.9357 Å at 100 K. The data were autoprocessed, indexed and integrated using xdsme v0.6.6⁵⁴ in space group P2₁2₁2 for the substrate-free structure and P2₁ for the substrate-bound structures. Data were scaled and merged using Aimless v0.7.8⁵⁵ in CCP4 v8.0⁵⁶ and truncated to 1.98 Å, 1.26 Å, and 1.12 Å for the substrate-free, syringol-bound and 4-methylsyringol-bound structures, respectively. Molecular replacement was performed using Phaser-MR v2.7.16⁵⁷ with an Alphafold2 v2.3.1⁵⁸ model of SyoA made using ColabFold v1.5.1⁵⁹ as the initial search model for the substrate-free structure, which was subsequently used to phase the syringol-bound model. The refined syringol-bound structure was then used as the search model to phase the 4-methylsyringol-bound data. The models were refined with Phenix.refine v1.20.1_4487 and built and improved in Coot v0.9.8.3⁶⁰ over multiple iterations. Data collection and refinement statistics are summarized in Supplementary Table 1. Molecular graphics, structural alignments and RMSD calculations were performed with UCSF ChimeraX v1.6.1⁶¹.

Substrate binding analysis and dissociation constants

To determine substrate binding, SyoA or GcoA was diluted to ~3 μM in 50 mM Tris pH 7.5 in 500 μL and substrate added in 1 μL aliquots from a 100 mM stock in DMSO. The absorbance was measured between 600–250 nm on a Cary 60 UV-Vis Spectrophotometer (Agilent Technologies) until no further shift was observed.

To determine substrate binding affinity (K_D), SyoA was diluted to ~2–8 μM in 50 mM potassium phosphate buffer pH 7.4 in a volume of 2.5 mL. Aliquots (0.5–5 μL) of 1, 10, or 100 mM substrate stock solutions in DMSO were added using a 5 μL Hamilton syringe. The difference spectrum was recorded from 300–600 nm. Aliquots of substrate were added until no further spectral shift occurred and no more than 10 μL of each stock solution was added to avoid diluting the enzyme. Titrations were performed in triplicate. Binding curve graphing analysis was performed with GraphPad Prism 10. The peak-to-trough absorbance difference, ΔA (A_peak – A_trough), was then plotted against substrate concentration. The ΔA was calculated using A_{390-419 nm} for syringol and 4-methylsyringol, A_{500-419 nm} for 4-allylsyringol and A_{500-570 nm} for syringaldehyde. To obtain the dissociation constant the data were fitted to the hyperbolic (Michaelis-Menten) equation (Equation 1), where K_D denotes the binding constant, [S] the substrate concentration, ΔA the peak-to-trough ratio, and ΔA_max the maximum peak-to-trough absorbance.

$$\Delta A=\frac{{\Delta A}_{\max }\times [S]}{{K}_{{{{\rm{D}}}}}+[S]}$$

(1)

In vitro NADH/O₂-driven reactions

In vitro NADH/O₂-driven reactions were performed at 30 °C in a total volume of 600–1200 μL. Reactions were performed in a quartz cuvette (with a 1 cm path length) and contained 1 μM SyoA or 0.5 μM GcoA, 1 μM SyoB or GcoB, and 100 μg mL⁻¹ catalase (from bovine liver) in oxygenated Tris buffer (50 mM, pH 7.5). This mixture was used to blank the spectrophotometer. NADH was added to a concentration of ∼320 μM and the background rate of NADH air oxidation was measured spectrophotometrically at 340 nm (ε340 nm = 6.22 mM^–1 cm^–1). The reaction was initiated by addition of 1 mM substrate (from a 100 mM stock solution in DMSO) and NADH oxidation was monitored at 340 nm for the first ∼10 min, and the reactions were then left at room temperature for ∼1 h to ensure that the NADH was completely consumed. Control reactions were also performed in which the P450 was omitted from the turnover mixture.

In vitro H₂O₂-driven reactions

In vitro H₂O₂ turnovers were performed at 30 °C in a total volume of 600 μL and contained 1 μM P450, 0.5–2 mM substrate from a 100 mM stock dissolved in DMSO and 10 mM H₂O₂ for the SyoA reactions and 20 mM H₂O₂ for the GcoA reactions in Tris buffer (50 mM, pH 7.5). Each reaction was performed in triplicate. Reactions were incubated for 2 mins at 30 °C prior to addition of H₂O₂ to initiate the reaction (H₂O₂ was added from a 200 mM stock freshly prepared before each experiment from a 30% w/w stock). All reactions were quenched at 60 min with 10 μL of 10 mg mL⁻¹ catalase. Control reactions without P450 were performed to assess stability of substrates and products with H₂O₂.

Formaldehyde assay

Formaldehyde production was measured using a colorimetric Purpald assay⁶². To determine the concentration of formaldehyde, 120 μL of the in vitro turnover mixture was mixed with 48 μL of 32 mM Purpald (dissolved in 2 M NaOH) and 432 μL 50 mM Tris, pH 7.5. The reaction was allowed to develop at room temperature by shaking for 60 min. The absorbance at 550 nm was recorded and the concentration of formaldehyde calculated using a calibration curve made with 0–400 μM formaldehyde. Reactions excluding the Purpald reagent were used as a baseline and the concentration of formaldehyde normalized against controls reactions without P450.

Product analysis by HPLC

Analytical High Performance Liquid Chromatography (HPLC) was performed on a Shimadzu LC-20AD equipped with a Phenomenex Kinetex 5 u XB-C18 100 Å column (250 mm×4.6 mm, 5 μM), SIL-20A autosampler, CTO-20A, SPD-20A UV detector, and CBM-20Alite communications module. Each sample and standard were injected at a volume of 20 μL. A gradient of 20–95% MeCN in water (with 0.1% TFA) over 30 min was used to elute the samples at a rate of 1 mL min⁻¹ and the eluate was monitored at 254 nm.

To prepare turnover mixtures for HPLC, 132 μL of the in vitro turnover mixture was mixed with 66 μL of MeCN and 2 μL of internal standard (IS, 10 mM 9-hydroxyfluorene in EtOH) and centrifuged at 15,800 × g for 3 mins to remove particulate matter. Calibration curves were constructed to quantify the product where available. Solutions of products with concentrations of 20, 50, 100, 200, and 500 μM were prepared for HPLC analysis in the same way as the turnovers. A plot of product peak area/IS peak area versus product concentration was then constructed.

Product analysis by GC-MS

In vitro H₂O₂ turnovers were performed as described above in 600 µL and analytes extracted once with 300 µL ethyl acetate. Samples were thoroughly mixed for 30 s by vortex. After phase separation, 180 µL of the organic layer was transferred to vials and analyzed by gas chromatography-mass spectrometry (GC-MS) using a Shimadzu GC-2010 equipped with a QP2010S GC-MS detector. 1 µL of sample was injected for analysis using a splitting ratio of 20. Separation of the analytes was done using a DB-5MS UI column (30 mm × 0.25 mm × 0.25 μm) at a flowrate of 1.4 mL min⁻¹ and the following temperature settings: Initial temperature of 100 °C held for 2 min, increased to 270 °C at 10 °C min⁻¹ and held at 270 °C for 6 min. The injection inlet temperature was 230 °C, ion source temperature 250 °C.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The atomic coordinates and structure factors generated in this study have been deposited in the Protein Data Bank under accession codes 8u09 SyoA; 8u19 syringol-bound SyoA, and 8u1i 4-methylsyringol-bound SyoA; structural comparisons were made to 5ncb guaiacol-bound GcoA. Source data including spectrophotometric data, HPLC data, per residue B-factors, RMSD measurements, binding affinity calculations, and absorbance data generated in this study are provided with this paper as a Source Data file. Source data are provided with this paper.

References

Wolf, M. E., Hinchen, D. J., DuBois, J. L., McGeehan, J. E. & Eltis, L. D. Cytochromes P450 in the biocatalytic valorization of lignin. Curr. Opin. Biotechnol. 73, 43–50 (2022).
Article CAS PubMed Google Scholar
Mallinson, S. J. B. et al. A promiscuous cytochrome P450 aromatic O-demethylase for lignin bioconversion. Nat. Commun. 9, 2487 (2018).
Article ADS PubMed PubMed Central MATH Google Scholar
Fetherolf, M. M. et al. Characterization of alkylguaiacol-degrading cytochromes P450 for the biocatalytic valorization of lignin. Proc. Natl Acad. Sci. USA 117, 25771–25778 (2020).
Article ADS CAS PubMed PubMed Central Google Scholar
Harlington, A. C., Shearwin, K. E., Bell, S. G. & Whelan, F. Efficient O-demethylation of lignin monoaromatics using the peroxygenase activity of cytochrome P450 enzymes. Chem. Commun. 58, 13321–13324 (2022).
Article CAS Google Scholar
Ralph, J., Lapierre, C. & Boerjan, W. Lignin structure and its engineering. Curr. Opin. Biotechnol. 56, 240–249 (2019).
Article CAS PubMed MATH Google Scholar
Becker, J. & Wittmann, C. A field of dreams: Lignin valorization into chemicals, materials, fuels, and health-care products. Biotechnol. Adv. 37, 107360 (2019).
Article CAS PubMed MATH Google Scholar
Abe, T., Masai, E., Miyauchi, K., Katayama, Y. & Fukuda, M. A tetrahydrofolate-dependent O-demethylase, LigM, is crucial for catabolism of vanillate and syringate in Sphingomonas paucimobilis SYK-6. J. Bacteriol. 187, 2030–2037 (2005).
Article CAS PubMed PubMed Central Google Scholar
Masai, E. et al. A novel tetrahydrofolate-dependent O-demethylase gene is essential for growth of Sphingomonas paucimobilis SYK-6 with syringate. J. Bacteriol. 186, 2757–2765 (2004).
Article CAS PubMed PubMed Central MATH Google Scholar
Perez, J. M. et al. Redundancy in aromatic O-demethylation and ring-opening reactions in Novosphingobium aromaticivorans and their impact in the metabolism of plant-derived phenolics. Appl. Environ. Microbiol 87, e02794–20 (2021).
Article CAS PubMed PubMed Central Google Scholar
Notonier, S. et al. Metabolism of syringyl lignin-derived compounds in Pseudomonas putida enables convergent production of 2-pyrone-4,6-dicarboxylic acid. Metab. Eng. 65, 111–122 (2021).
Article CAS PubMed Google Scholar
Machovina, M. M. et al. Enabling microbial syringol conversion through structure-guided protein engineering. Proc. Natl Acad. Sci. USA 116, 13970–13976 (2019).
Article ADS CAS PubMed PubMed Central Google Scholar
Jiang, Y. et al. Regioselective aromatic O-demethylation with an artificial P450BM3 peroxygenase system. Catal. Sci. Technol. 10, 1219–1223 (2020).
Article CAS MATH Google Scholar
Guengerich, F. P. Mechanisms of cytochrome P450-catalyzed oxidations. ACS Catal. 8, 10964–10976 (2018).
Article CAS PubMed PubMed Central MATH Google Scholar
Huang, X. & Groves, J. T. Oxygen activation and radical transformations in heme proteins and metalloporphyrins. Chem. Rev. 118, 2491–2553 (2018).
Article CAS PubMed MATH Google Scholar
Rittle, J. & Green, M. T. Cytochrome P450 compound I: capture, characterization, and C-H bond activation kinetics. Science 330, 933–937 (2010).
Article ADS CAS PubMed Google Scholar
Krest, C. M. et al. Reactive intermediates in cytochrome P450 catalysis. J. Biol. Chem. 288, 17074–17081 (2013).
Article CAS PubMed PubMed Central Google Scholar
Imai, M. et al. Uncoupling of the cytochrome P-450cam monooxygenase reaction by a single mutation, threonine-252 to alanine or valine: A possible role of the hydroxy amino acid in oxygen activation. Proc. Natl Acad. Sci. USA 86, 7823–7827 (1989).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Raag, R. et al. Crystal structure of the cytochrome P-450CAM active site mutant Thr252. Ala. Biochemistry 30, 11420–11429 (1991).
Article CAS PubMed MATH Google Scholar
Kimata, Y., Shimada, H., Hirose, T. & Ishimura, Y. Role of Thr-252 in cytochrome P450cam: a study with unnatural amino acid mutagenesis. Biochem. Biophys. Res. Commun. 208, 96–102 (1995).
Article CAS PubMed Google Scholar
Vidakovic, M., Sligar, S. G., Li, H. & Poulos, T. L. Understanding the role of the essential Asp251 in cytochrome P450cam using site-directed mutagenesis, crystallography, and kinetic solvent isotope effect. Biochemistry 37, 9211–9219 (1998).
Article CAS PubMed Google Scholar
Coleman, T. et al. Structural insights into the role of the acid-alcohol pair of residues required for dioxygen activation in cytochrome P450 enzymes. J. Biol. Inorg. Chem. 25, 583–596 (2020).
Article CAS PubMed MATH Google Scholar
Shoji, O. & Watanabe, Y. Peroxygenase reactions catalyzed by cytochromes P450. J. Biol. Inorg. Chem. 19, 529–539 (2014).
Article CAS PubMed MATH Google Scholar
Hrycay, E. G. & Bandiera, S. M. “Monooxygenase, peroxidase and peroxygenase properties and reaction mechanisms of cytochrome P450 enzymes” in Monooxygenase, Peroxidase and Peroxygenase Properties and Mechanisms of Cytochrome P450. Advances in Experimental Medicine and Biology (eds Hrycay, E. G. & Bandiera, S. M.) 1–61 (Springer, 2015).
Albertolle, M. E. & Guengerich, P. F. The relationships between cytochromes P450 and H₂O₂: production, reaction, and inhibition. J. Inorg. Biochem 186, 228–234 (2018).
Article CAS PubMed PubMed Central Google Scholar
Matsunaga, I., Ueda, A., Fujiwara, N., Sumimoto, T. & Ichihara, K. Characterization of the ybdT gene product of Bacillus subtilis: novel fatty acid β-hydroxylating cytochrome P450. Lipids 34, 841–846 (1999).
Article CAS PubMed Google Scholar
Belcher, J. et al. Structure and biochemical properties of the alkene producing cytochrome P450 OleTJE (CYP152l1) from the Jeotgalicoccus sp. 8456 bacterium. J. Biol. Chem. 289, 6535–6550 (2014).
Article CAS PubMed PubMed Central MATH Google Scholar
Munro, A. W., McLean, K. J., Grant, J. L. & Makris, T. M. Structure and function of the cytochrome P450 peroxygenase enzymes. Biochem. Soc. Trans. 46, 183–196 (2018).
Article CAS PubMed PubMed Central MATH Google Scholar
Ebrecht, A. C., Smit, M. S. & Opperman, D. J. Natural alternative heme-environments allow efficient peroxygenase activity by cytochrome P450 monooxygenases. Catal. Sci. Technol. 13, 6264–6273 (2023).
Article CAS Google Scholar
Hofrichter, M., Kellner, H., Pecyna, M. J. & Ullrich, R. Fungal unspecific peroxygenases: heme-thiolate proteins that combine peroxidase and cytochrome P450 properties. Adv. Exp. Med. Biol. 851, 341–368 (2015).
Article CAS PubMed Google Scholar
Correll, C. C., Batie, C. J., Ballou, D. P. & Ludwig, M. L. Phthalate dioxygenase reductase: a modular structure for electron transfer from pyridine nucleotides to [2Fe-2S]. Science 258, 1604–1610 (1992).
Article ADS CAS PubMed Google Scholar
Dakin, H. D. The oxidation of hydroxy derivatives of benzaldehyde, acetophenone and related substances. Am. Chem. J. 42, 477–498 (1909).
MATH Google Scholar
Nagano, S. & Poulos, T. L. Crystallographic study on the dioxygen complex of wild-type and mutant cytochrome P450cam. Implications for the dioxygen activation mechanism. J. Biol. Chem. 280, 31659–31663 (2005).
Article CAS PubMed Google Scholar
Harth, F. M. et al. Selective demethylation reactions of biomass-derived aromatic ether polymers for bio-based lignin chemicals. Green. Chem. 25, 10117–10143 (2023).
Article CAS MATH Google Scholar
Winkler, C. K., Schrittwieser, J. H. & Kroutil, W. Power of biocatalysis for organic synthesis. ACS Cent. Sci. 7, 55–71 (2021).
Article CAS PubMed PubMed Central MATH Google Scholar
Zhou, N., Thilakarathna, W. W., He, Q. S. & Rupasinghe, H. V. A review: depolymerization of lignin to generate high-value bio-products: opportunities, challenges, and prospects. Front. Energy Res. 9, 758744 (2022).
Lee, S., Kang, M., Bae, J. H., Sohn, J. H. & Sung, B. H. Bacterial valorization of lignin: strains, enzymes, conversion pathways, biosensors, and perspectives. Front. Bioeng. Biotechnol. 7, 209 (2019).
Article PubMed PubMed Central MATH Google Scholar
Lu, X. & Gu, X. A review on lignin pyrolysis: pyrolytic behavior, mechanism, and relevant upgrading for improving process efficiency. Biotechnol. Biofuels Bioprod. 15, 106 (2022).
Article CAS PubMed PubMed Central MATH Google Scholar
Lee, Y. T., Wilson, R. F., Rupniewski, I. & Goodin, D. B. P450cam visits an open conformation in the absence of substrate. Biochem 49, 3412–3419 (2010).
Article CAS MATH Google Scholar
Santos, S. F. G., Bommareddy, R. R., Black, G. W., Singh, W. & Huang, M. The substrate specificity in the O-demethylation of 4-alkylguaiacols by cytochrome P450 AgcAP450. Catal. Sci. Technol. 13, 2070–2079 (2023).
Article CAS Google Scholar
Hall, E. A., Sarkar, M. R. & Bell, S. G. The selective oxidation of substituted aromatic hydrocarbons and the observation of uncoupling via redox cycling during naphthalene oxidation by the CYP101B1 system. Catal. Sci. Technol. 7, 1537–1548 (2017).
Article CAS MATH Google Scholar
Zhao, L. et al. Biological degradation of lignin: a critical review on progress and perspectives. Ind., Crop. Prod. 188, 115715 (2022).
Article CAS MATH Google Scholar
Erickson, E. et al. Critical enzyme reactions in aromatic catabolism for microbial lignin conversion. Nat. Catal. 5, 86–98 (2022).
Article CAS MATH Google Scholar
Sun, S. et al. Lytic polysaccharide monooxygenase synergized with lignin-degrading enzymes for efficient lignin degradation. iScience 26, 107870 (2023).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Lahtinen, M., Kruus, K., Heinonen, P. & Sipilä, J. On the reactions of two fungal laccases differing in their redox potential with lignin model compounds: products and their rate of formation. J. Agric. Food Chem. 57, 8357–8365 (2009).
Article CAS PubMed Google Scholar
Kim, J., Nguyen, T. V. T., Kim, Y. H., Hollmann, F. & Park, C. B. Lignin as a multifunctional photocatalyst for solar-powered biocatalytic oxyfunctionalization of C–H bonds. Nat. Synth. 1, 217–226 (2022).
Article ADS CAS MATH Google Scholar
Kommedal, E. G. et al. Visible light-exposed lignin facilitates cellulose solubilization by lytic polysaccharide monooxygenases. Nat. Commun. 14, 1063 (2023).
Article ADS CAS PubMed PubMed Central Google Scholar
Li, F. et al. Microbial lignin valorization through depolymerization to aromatics conversion. Trends Biotechnol. 40, 1469–1487 (2022).
Article CAS PubMed MATH Google Scholar
Gee, A. R. et al. Efficient biocatalytic C–H bond oxidation: an engineered heme-thiolate peroxygenase from a thermostable cytochrome P450. Chem. Commun. 59, 13486–13489 (2023).
Article CAS Google Scholar
Podgorski, M. N. et al. Engineering peroxygenase activity into cytochrome P450 monooxygenases through modification of the oxygen binding region. ACS Catal. 14, 7426–7443 (2024).
Article CAS MATH Google Scholar
Gibson, D. G. et al. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat. Methods 6, 343–345 (2009).
Article CAS PubMed MATH Google Scholar
De Marco, A., Vigh, L., Diamant, S. & Goloubinoff, P. Native folding of aggregation-prone recombinant proteins in Escherichia coli by osmolytes, plasmid- or benzyl alcohol-overexpressed molecular chaperones. Cell Stress Chaperones 10, 329–339 (2005).
Article PubMed PubMed Central Google Scholar
Cowieson, N. P. et al. MX1: a bending-magnet crystallography beamline serving both chemical and macromolecular crystallography communities at the Australian Synchrotron. J. Synchrotron Radiat. 22, 187–190 (2015).
Article CAS PubMed PubMed Central Google Scholar
Aragão, D. et al. MX2: a high-flux undulator microfocus beamline serving both the chemical and macromolecular crystallography communities at the Australian Synchrotron. J. Synchrotron Radiat. 25, 885–891 (2018).
Article PubMed PubMed Central Google Scholar
Legrand, P. XDSME: XDS made easier. GitHub repository. accessed 16 January 2024. Available at: https://github.com/legrandp/xdsme (2017).
Evans, P. R. & Murshudov, G. N. How good are my data and what is the resolution? Acta Crystallogr. D. Biol. Crystallogr. 69, 1204–1214 (2013).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Agirre, J. et al. The CCP4 suite: integrative software for macromolecular crystallography. Acta Crystallogr. D. Struct. Biol. 79, 449–461 (2023).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Jumper, J. et al. Highly accurate protein structure prediction with AlphaFold. Nature 596, 583–589 (2021).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Mirdita, M. et al. ColabFold: making protein folding accessible to all. Nat. Methods 19, 679–682 (2022).
Article CAS PubMed PubMed Central MATH Google Scholar
Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D. Biol. Crystallogr. 66, 486–501 (2010).
Article ADS CAS PubMed PubMed Central MATH Google Scholar
Meng, E. C. et al. UCSF ChimeraX: tools for structure building and analysis. Prot. Sci. 32, 4792 (2023).
Article MATH Google Scholar
Quesenberry, M. S. & Lee, Y. C. A rapid formaldehyde assay using purpald reagent: application under periodation conditions. Anal. Biochem. 234, 50–55 (1996).
Article CAS PubMed Google Scholar

Download references

Acknowledgements

This work was funded, in part, through Australian Research Council grants DP200102411 (to S.G.B. and others) and DP230103062 (to F.W. and K.E.S.). This research was undertaken on the MX1 and MX2 beamlines at the Australian Synchrotron, part of Australian Nuclear Science and Technology Organization, and made use of the ACRF detector on MX2. We acknowledge financial support from the Australian Synchrotron to use this facility. A.C.H. was supported by Wine Australia, with levies from Australia’s grapegrowers and winemakers and matching funds from the Australian Government. A.C.H. and T.D. were supported by Research Training Program (RTP) scholarships from the Australian government. F.W. was supported by the Ramsay Fellowship in Applied Science and Microscopy Australia (ROR: 042mm0k03). We acknowledge that this research was conducted on the traditional Country of the Kaurna people and the Boonwurrung/Bunurong and Wurundjeri people of the Kulin Nation and pay our respects to Elders past and present.

Author information

Authors and Affiliations

Department of Molecular and Biomedical Science, University of Adelaide, Adelaide, SA, Australia
Alix C. Harlington, Keith E. Shearwin & Fiona Whelan
Department of Chemistry, University of Adelaide, Adelaide, SA, Australia
Tuhin Das & Stephen G. Bell
Adelaide Microscopy, University of Adelaide, Adelaide, SA, Australia
Fiona Whelan

Authors

Alix C. Harlington
View author publications
Search author on:PubMed Google Scholar
Tuhin Das
View author publications
Search author on:PubMed Google Scholar
Keith E. Shearwin
View author publications
Search author on:PubMed Google Scholar
Stephen G. Bell
View author publications
Search author on:PubMed Google Scholar
Fiona Whelan
View author publications
Search author on:PubMed Google Scholar

Contributions

A.C.H., T.D., K.E.S., S.G.B., and F.W. designed research; A.C.H., T.D., K.E.S., S.G.B., and F.W. performed research; A.C.H., T.D., S.G.B., and F.W. analyzed data; A.C.H., K.E.S., S.G.B., and F.W. wrote the paper.

Corresponding authors

Correspondence to Stephen G. Bell or Fiona Whelan.

Ethics declarations

Competing interests

The authors declare no competing interests.

Peer review

Peer review information

Nature Communications thanks Jing-Ke Weng and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Supplementary Information

Reporting Summary

Transparent Peer Review file

Source data

Source Data

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.

Reprints and permissions

About this article

Cite this article

Harlington, A.C., Das, T., Shearwin, K.E. et al. Structural insights into S-lignin O-demethylation via a rare class of heme peroxygenase enzymes. Nat Commun 16, 1815 (2025). https://doi.org/10.1038/s41467-025-57129-6

Download citation

Received: 17 July 2024
Accepted: 05 February 2025
Published: 20 February 2025
DOI: https://doi.org/10.1038/s41467-025-57129-6