Polysaccharide oxidation by lytic polysaccharide monooxygenase is enhanced by engineered cellobiose dehydrogenase

Abstract

The catalytic function of lytic polysaccharide monooxygenases (LPMOs) to cleave and decrystallize recalcitrant polysaccharides put these enzymes in the spotlight of fundamental and applied research. Here we demonstrate that the demand of LPMO for an electron donor and an oxygen species as cosubstrate can be fulfilled by a single auxiliary enzyme: an engineered fungal cellobiose dehydrogenase (CDH) with increased oxidase activity. The engineered CDH was about 30 times more efficient in driving the LPMO reaction due to its 27 time increased production of H₂ O₂ acting as a cosubstrate for LPMO. Transient kinetic measurements confirmed that intra- and intermolecular electron transfer rates of the engineered CDH were similar to the wild-type CDH, meaning that the mutations had not compromised CDH's role as an electron donor. These results support the notion of H₂ O₂ -driven LPMO activity and shed new light on the role of CDH in activating LPMOs. Importantly, the results also demonstrate that the use of the engineered CDH results in fast and steady LPMO reactions with CDH-generated H₂ O₂ as a cosubstrate, which may provide new opportunities to employ LPMOs in biomass hydrolysis to generate fuels and chemicals.

Keywords: cellobiose dehydrogenase; cellulose degradation; copper monooxygenase; hydrogen peroxide; lytic polysaccharide monooxygenase.

Figure 1

The active site of ChCDH in complex with the inhibitor cellobionolactam (CBLM, pdb: http://www.rcsb.org/pdb/search/structidSearch.do?structureId=4QI5). Shown is the FAD‐cofactor (orange) with the indicated C4a and N5 atoms and the catalytic base H701 (blue). The positions N700 (magenta) and N748 (green) of the wild‐type ChCDH were replaced in this study by serine and glycine, respectively.

Figure 2

Product formation by SmLPMO10A. Chitobionic acid concentration after incubation of β‐chitin with 1 µm SmLPMO10A, 15 mm lactose and 10 different concentrations of ChCDH (blue) or CDH_oxy+ (red) after 1 h (A) or 24 h (B). Panel (C) shows the time‐dependent formation of chitobionic acid by SmLPMO10A and (D) lactobionic acid by CDH_oxy+ in reactions containing 1 µm LPMO and five different concentrations of CDH_oxy+. All error bars show ± S.D. (n = 3).

Figure 3

Oxidized sites generated during chitin degradation by SmLPMO10A. Panel (A) shows a time‐independent plot of the degree of solubilization of oxidized products (i.e., the percentage of products obtained in the soluble fraction compared to the total number of oxidized sites) as a function of the quantity of detected solubilized oxidized products. The total number of oxidized sites was determined in reactions containing 10 g·L⁻¹ β‐chitin and 1 µm SmLPMO10A with varying concentrations of CDH_oxy+ (31.25 or 125 nm) and 15 mm lactose. Samples were taken at different stages of the reaction (15–180 min). The samples were diluted twice and heat‐inactivated (100 °C, 15 min). Half of the reaction was filtrated and 1 µm SmGH20 chitobiase was added for quantitation of solubilized products (i.e., the standard procedure for quantitation of solubilized oxidized sites). To the other half of the reaction, a cocktail of chitinases (containing 2 µm SmChi18A, 2 µm SmChi18C and 1 µm SmGH20) was added followed by incubation for 20 h at 37 °C until the chitin was fully degraded. The resulting chitobionic acid concentration represents the total amount of oxidized sites. Rates derived from the initial phases of the progress curves in Fig. 2C,D only included time points where the solubilization was < 200 µm which means that the total number of oxidized sites is underestimated by ~ 50%. Panel (B) shows the LPMO rate without and with correction for this underestimation. The corrected curve shows that 1 molecule of lactose converted by CDH_oxy+ leads to approximately 0.8 molecules of chitobionic acid being produced by SmLPMO10A.

Figure 4

Product formation by NcLPMO9C. HPAEC‐PAD chromatograms show C4‐oxidized products generated in reactions containing 1 µm NcLPMO9C in the presence of 62.5 nm (A) or 1500 nm ChCDH (B) and 62.5 nm (C) or 1500 nm CDH_oxy+ (D). All reactions contained 5 g·L⁻¹ PASC and 15 mm lactose and were incubated in 50 mm potassium phosphate buffer, pH 6.0, for up to 24 h in an Eppendorf thermomixer (Hamburg, Germany) set to 30 °C and 800 r.p.m. Samples were heat‐inactivated (15 min, 100 °C) and filtered (0.22 µm) prior to analysis. The bigger black arrows indicate the Glc4GemGlc C4‐oxidized product, whereas the smaller black arrows, seen in panel (B–D), show Glc₂Glc1A peaks that are formed as a result of CDH oxidation of the native cellotriose that is formed by the LPMO upon cleavage of shorter cello‐oligosaccharides. Peak annotations are based on earlier studies 41, 42. Note that the elution times vary slightly between different runs. Double (C1/C4) oxidized products (peaks eluting after ~ 40 min) dominate since CDH oxidizes the reducing end of the C4‐oxidized products formed by the LPMO. Note that C4‐oxidized products are unstable, which explains why some peaks become smaller over time in panel (D).

Figure 5

Electron transfer between redox centers. Intramolecular electron transfer in ChCDH (A) and CDH_oxy+ (B) from FAD, in the DH domain, to heme b, in the cytochrome domain, at the indicated cellobiose concentrations. Heme b reduction was followed at 563 nm (α‐band). The concentration of CDH after mixing was 1.5 µm. Prior to all measurements, substrate and enzyme solutions were carefully degassed by applying alternating cycles of vacuum and nitrogen pressure to avoid interference with atmospheric oxygen. All measurements were carried out at 30 °C in 50 mm sodium phosphate buffer, pH 6.0. The insets show the cellobiose concentration‐dependent electron transfer rates. Panels (C) and (D) show raw traces for heme b reoxidation by SmLPMO10A in the absence (catalase added, C) or presence (no catalase added, D) of H₂O₂ using 0 µm (gray), 15 µm (green), 30 µm (blue) or 60 µm (red) LPMO. (E) Electron transfer from reduced heme b to LPMO in the presence (red) or absence (blue) of approximately 250 µm H₂O₂ formed by CDH during prereduction. All traces are the average of triplicate measurements. Error bars show ± S.D (n = 3).

Figure 6

LPMO activity driven by CDH without the cytochrome domain. (A) Time courses of chitobionic acid production by 1 µm SmLPMO10A in combination with CDH variants. (B) Quantitation of LPMO products generated in reactions containing 0.25 mm FAD or 0.1 mm ascorbic acid (AscA) in the presence or absence of 750 nm ChDH or 31.25 nm DH_oxy+. Relative activities show the enhancement upon addition of FAD or AscA (based on product levels after 24 h). The error bars show ± S.D (n = 3). Lower panels: Reduction of SmLPMO10A by different CDH variants under anaerobic or aerobic conditions. All reactions contained 2 µm SmLPMO10A and 0.25 µm ChCDH (C); 0.25 µm ChDH (D); or 31.25 nm DH_oxy+ (E). Reactions were initiated by addition of lactose (1 mm final concentration, black arrow) and were performed in duplicate (both traces are displayed). The variation in fluorescence is relative to the maximum increase in fluorescence measured for a control reaction (black line) in which lactose was replaced by 10 µm AscA, which represents a 100% reduction of SmLPMO10A. The " //" labels in panel (D) and (E) indicate when the reactions were no longer under anaerobic conditions.

Figure 7

Stoichiometry of LPMO reduction with FADH₂. Titration of FADH₂ with NcLPMO9C (A) or SmLPMO10A (B). Oxidized FAD (60 µm) was approximately 70% reduced with sodium dithionite. The FADH₂ was reoxidized with LPMO by adding aliquots of 3 µL (NcLPMO9C; 515 µm) or 1.5 µL (SmLPMO10A; 906 µm) to the cuvette. Absorption spectra were recorded with an Agilent 8453 UV/VIS‐Spectrometer featuring a diode array detector. Blank reactions (buffer titrated to FADH₂ or LPMO) were subtracted. The concentration of FAD and LPMOs was determined based on their molar absorption coefficients (FAD ε₄₅₀ = 11.3 mm ⁻¹·cm⁻¹; NcLPMO9C ε₂₈₀ = 46.91 mm ⁻¹·cm⁻¹; SmLPMO10A ε₂₈₀ = 35.20 mm ⁻¹·cm⁻¹). All reactions were carried out at 23 °C in an anaerobic glove box (Whitley DG250, Don Whitley Scientific) flushed with a nitrogen/hydrogen mixture. The data show that, for both LPMOs, about two molecules of enzyme were required to reoxidize 1 molecule of FADH₂.