Discovery of fungal oligosaccharide-oxidising flavo-enzymes with previously unknown substrates, redox-activity profiles and interplay with LPMOs

Abstract

Oxidative plant cell-wall processing enzymes are of great importance in biology and biotechnology. Yet, our insight into the functional interplay amongst such oxidative enzymes remains limited. Here, a phylogenetic analysis of the auxiliary activity 7 family (AA7), currently harbouring oligosaccharide flavo-oxidases, reveals a striking abundance of AA7-genes in phytopathogenic fungi and Oomycetes. Expression of five fungal enzymes, including three from unexplored clades, expands the AA7-substrate range and unveils a cellooligosaccharide dehydrogenase activity, previously unknown within AA7. Sequence and structural analyses identify unique signatures distinguishing the strict dehydrogenase clade from canonical AA7 oxidases. The discovered dehydrogenase directly is able to transfer electrons to an AA9 lytic polysaccharide monooxygenase (LPMO) and fuel cellulose degradation by LPMOs without exogenous reductants. The expansion of redox-profiles and substrate range highlights the functional diversity within AA7 and sets the stage for harnessing AA7 dehydrogenases to fine-tune LPMO activity in biotechnological conversion of plant feedstocks.

Fig. 1. Phylogenetic analysis of AA7-like sequences.

a The phylogenetic tree is based on 1927 sequences. Biochemically characterized enzymes are coloured according to clade, with green circles indicating enzymes from the present study. The PDB entries (in square brackets) are given for available enzyme structures. Clade Va harbours the canonical previously described oligosaccharide oxidases: SsGOOX from Sarocladium strictum active on cellooligosaccharides, MnLaO from Microdochium nivale active on lactose, TtXylO from Thermothelomyces thermophilus active on xylooligosaccharides and FgChitO from Fusarium graminearum active on chitooligosaccharides. Clade I contains characterised plant enzymes^, that are not assigned in AA7 including the oligogalacturonide oxidase (AtOGOX1) from Arabidopsis thaliana^,,. b Clade-wise taxonomic distribution of putative AA7 sequences. c Clade-wise conservation percentage of the histidine and cysteine FAD-cofactor tethering residues. The number of sequences within each clade is indicated above each bar. The accessions of the sequences in the tree are provided in the Source Data file.

Fig. 2. Spectral properties and time-course NMR analysis of the AA7 cellooligosaccharide dehydrogenase FgCelDH7C.

a Spectral comparison of the flavin absorbance of FgCelDH7C and the oxidases FgChi7B as well as MoChi7A, all at 20 µM. The data are means ± standard deviations (n = 3 independent experiments). b and c Show the time resolved in situ ¹H NMR analyses of the oxidation of cellobiose by FgCelDH7C in the presence (b) or absence (c) of 1.3 mM 2,6-dichlorophenolindophenol (DCIP) as an electron acceptor, respectively. The presented spectra in panels b and c are from a single experiment (n = 1). Source data are provided as a Source Data file.

Fig. 3. Active site signatures of AA7 oligosaccharide oxidases versus dehydrogenases.

The active sites of the clade IIa discovered dehydrogenase FgCelDH7C (PDB: 6YJI) and the canonical clade Va cellooligosaccharide oxidase SsGOOX (in complex with 5-amino-5-deoxy-cellobiono-1,5-lactam ABL, PDB: 2AXR) are shown. a The FAD-tethering histidine in addition to an aromatic cluster comprising the tyrosine base catalyst, a phenylalanine/tyrosine and the substrate-stacking aromatic residue are conserved features in fungal oligosaccharide dehydrogenases and oxidases. b Sequence logos of patches spanning the structurally similar active site residues shown in (a). c Active site differences between clades IIa and Va. d Sequence logos of patches spanning the structurally divergent active site residues shown in c from clades IIa and Va. The amino acid numbering of the deposited protein sequences is used in the figure.

Fig. 4. Analysis of AA7-LPMO interplay in cellulose degradation.

Reactions were performed on Avicel using FgCelDH7C and PaLPMO9H with subsequent ionic chromatography (HPAEC-PAD) analysis. a A representative chromatogram part showing C1-C4 double-oxidised species in the cellulose degradation assay including combinations of Avicel (5 mg mL⁻¹), FgCelDH7C (0.4 μM), PaLPMO9H (4 μM), PaCDHB (1.2 µM), cellotetraose (DP4, 0.8 mM) and ascorbate (Asc., 1 mM) as indicated in the figure. b Comparison of the cellulose degradation assay based on the cumulative area under the peaks of native (DP3, DP5 and DP6), C1 oxidised (except DP2 and DP4 which were added as substrates for the CDH and AA7, respectively) and C1-C4 double-oxidised cellooligosaccharides from the reactions in (a). The data in (a) and (b) (n = 3 independent reactions) were generated in NaOAc/NaOH buffer (50 mM, pH 5.2) at 35 °C. The bar plot in (b) shows the means of total peak area (n = 3 independent reactions, each shown as a white circle) with standard deviations. Source data are provided as a Source Data file.

Fig. 5. Mechanistic insights into the activation of LPMO by FgCelDH7C.

a, b Effect of horseradish peroxidase (HRP) and superoxide dismutase (SOD) on the interplay between PaLPMO9H (4 μM) and FgCelDH7C (0.41 μM) in Avicel degradation based on the total area of native, C1 oxidised (C1 ox) and double C1-C4 oxidised (C1-C4ox) cello-oligosaccharides as analysed by HPAEC-PAD. Controls prepared in the absence of FgCelDH7C (1) or in the absence of both SOD and FgCelDH7C (2). The assays were performed for 18 h at 35 °C and terminated using NaOH (0.1 M) prior to the HPAEC-PAD analyses. The total peak area (white circles, n = 3 independent experiments) are shown and the bar plots display the means with standard deviations. c X-band Electron Paramagnetic Resonance (EPR) spectra of PaLPMO9H−Cu(II) (100 μM) in the presence of cellotriose (DP3, 1 mM) before (blue line) and after (red line) addition of FgCelDH7C (AA7). d X-band EPR spectra of PaLPMO9H−Cu(II) (20 μM) in buffer (blue line), in the presence of FgCelDH7C (AA7) pre-reduced with dithionite (red line), or directly fully reduced with dithionite (10 eq., green line). All EPR solutions and experiments were performed under anaerobic conditions. Samples were in 50 mM NaOAc, pH 5.2 and spectra were recorded at 50 K with a 4 mW microwave power and a 30 Gauss modulation amplitude. The data in (c) and (d) are based on a single experiment (n = 1). Source data for the a and b panels are provided as a Source Data file.

Fig. 6. Oxygen-binding cavity in AA7 oxidoreductases and a schematic model for AA7-LPMO interplay.

a, b Putative oxygen-binding cavity in typical AA7 oxidases and dehydrogenases, respectively. The substitution of conserved histidine and glycine in oxidases to serine (or other small residues) and valine in dehydrogenases, respectively, is observed. c Schematic model of the interplay of cellobiose dehydrogenase (CDH) versus the AA7 cellooligosaccharide dehydrogenase with LPMOs during cellulose degradation. Both dehydrogenase classes oxidise cellooligosaccharides to the corresponding lactones. The electrons harvested from this oxidation are stored in the FAD-cofactor and subsequently delivered directly to the LPMO in the case of AA7. The transfer of priming electrons in CDH proceeds first from the dehydrogenase domain to the cytochrome b haem domain in (closed form). A subsequent large conformational change to the open form is required to expose the haem domain to the LPMO active site for electron transfer. The dotted lines signify the low oxidase side-activity from both classes of dehydrogenases, which generates the H₂O₂ preferred co-substrate to fuel cellulose oxidative degradation by LPMOs. Low levels of H₂O₂ are also generated at the active site of free primed LPMOs, but this is left out from the figure for clarity.