A fast and sensitive activity assay for lytic polysaccharide monooxygenase
Lytic polysaccharide monooxygenases (LPMO) release a spectrum of cleavage products from their polymeric substrates cellulose, hemicellulose, or chitin. The correct identification and quantitation of these released products is the basis of MS/HPLC-based detection methods for LPMO activity. The duration, effort, and intricate analysis allow only specialized laboratories to measure LPMO activity in day-to-day work. A spectrophotometric assay will simplify the screening for LPMO in culture supernatants, help monitor recombinant LPMO expression and purification, and support enzyme characterization.
Based on a newly discovered peroxidase activity of LPMO, we propose a fast, robust, and sensitive spectrophotometric activity assay using 2,6-dimethoxyphenol (2,6-DMP) and H2O2. The fast enzymatic assay (300 s) consists of 1 mM 2,6-DMP as chromogenic substrate, 100 µM H2O2 as cosubstrate, and an adequate activity of LPMO in a suitable buffer. The high molar absorption coefficient of the formed product coerulignone (ε469 = 53,200 M−1 cm−1) makes the assay sensitive and allows reliable activity measurements of LPMO in concentrations of approx. 0.5–50 mg L−1.
KeywordsActivity assay 2,6-Dimethoxyphenol Hydrogen peroxide Biomass degradation Lytic polysaccharide monooxygenase Peroxidase activity
charged aerosol detection
degree of polymerization
electrospray ionization mass spectrometry
fluorescence resonance energy transfer
high performance anion exchange chromatography
hydrophilic interaction liquid chromatography
limit of blank
limit of detection
lytic polysaccharide monooxygenase
matrix-assisted laser desorption ionization-time of flight mass spectrometry
polysaccharide analysis by carbohydrate gel electrophoresis
phosphoric acid swollen cellulose
porous graphitized carbon
ultra high performance liquid chromatography
Lytic polysaccharide monooxygenases (LPMO; CAZy auxiliary activities AA9, AA10, AA11, and AA13 ; preliminary EC classification 1.14.99.B6) catalyze the oxidative depolymerization of diverse polymeric carbohydrates such as cellulose (AA9, AA10), hemicellulose (AA9, AA14 ), chitin (AA10, AA11), and starch (AA13). The polymeric nature of these substrates and the wide spectrum of released products make the assessment of LPMO activity difficult. Typically, LPMO and a polymeric substrate are incubated under defined assay conditions in vials for a defined period (up to 1 day) before a calibrated HPLC method is used for the quantitation of the products. If a new LPMO/substrate combination is measured, the reaction products have to be identified by HPLC standards or mass spectrometry. Protocols for the determination of reaction products were published for cellulose (HPAEC and MALDI-TOF [3, 4]; PGC, CAD and ESI-MS ), hemicellulose (HPAEC and ESI-MS ; HPAEC and MALDI-TOF, ), and chitin (UHPLC-HILIC coupled to MALDI-TOF, [8, 9], and RP-UHPLC-MS ). HPLC-MS measurements of LPMO activity give detailed insight into the formed reaction products, but are time consuming. The detection of reaction products is restricted to low molecular weight cleavage products (DP 1–6), which are soluble in the reaction buffer and the HPLC eluent, and for which HPLC standards are available. The quantitation of larger products that remain in the insoluble fraction can be achieved by total hydrolysis. By this way, C1-oxidized products have been quantified by HPLC, but the quantification of C4-oxidized products remains a struggle. Also, the most commonly used cellulosic substrate is phosphoric acid swollen cellulose (PASC), which is not a physiological substrate of LPMO. Nevertheless, if suitable reaction conditions, sampling time points and correct calibration data for the detected species are used, HPLC-based methods provide the most accurate results of physiological LPMO activity. NMR has also been used to follow the action of LPMO on polymeric substrates [4, 6, 11, 12], but its strength lies in the identification of reaction products and not in the measurement of enzymatic activity.
An activity measurement of LPMO on PASC, analyzed with polysaccharide analysis by carbohydrate gel electrophoresis (PACE), has been reported to yield mostly cellobiose and cellotriose . The PACE method allows the use of natural substrates and multiple samples and gives semi-quantitative results; however, the effort for preparation and measurement is much higher compared to a spectrophotometrical activity assay. It was also shown that the used C4-oxidizing LPMO is able to cleave cellotriose, which demonstrates that LPMOs can act on small oligosaccharides and perform successive depolymerization reactions. This feature of some LPMOs additionally makes an absolute quantitation of reaction products difficult. The use of derivatized cello-oligosaccharides as a substrate to follow cleavage by FRET quenching is far less affected by successive depolymerization. This method was used to determine kinetic constants of LPMO for a fluorescence-labeled cellotetraose  and is certainly qualified to measure homogeneous LPMO activity when the substrate becomes available. A microarray method to detect LPMO activity on hemicelluloses was established and gave semi-quantitative results . A microplate-based detection of insoluble C1-oxidized products of LPMO was achieved by fluorescence-labeling with 7-amino-1,3-naphthalene-disulfonic acid (ANDA). The method was verified by X-ray photoelectron spectroscopy . Finally, chromogenic polysaccharide hydrogel substrates were used to screen for the activity of different carbohydrate-active enzymes including LPMO. This semi-quantitative method can detect the cleavage of specific oligosaccharides by LPMO and can be used to screen for its activity on agar plates .
In contrast to these methods, which all detect the more or less natural activity of LPMO, stands a method which measures LPMO activity in a spectrophotometric/-fluorimetric assay based on the release of H2O2, which is a product of the uncoupling reaction of the activated copper–oxygen complex in the absence of substrate. The Amplex red assay is a coupled enzyme assay , which uses the formation of hydrogen peroxide by LPMO in the presence of oxygen and a reductant like ascorbic acid or cellobiose dehydrogenase (CDH) for the peroxidase-catalyzed oxidation of Amplex red to resorufin which can be detected spectrophotometrically (ε571 = 58,000 M−1 cm−1) or fluorimetrically (Ex = 569 nm/Em = 585 nm). This assay can be used to detect the production and purification of LPMOs; however, the uncoupling reaction is slow, the sensitivity is low, and the assay requires a high LPMO concentration (20–574 mg L−1) for reliable measurements. It is also affected by metal ions present in fermentation media, which requires re-buffering in spin columns or diafiltration. The Amplex red-based assay is most useful for quick activity measurements to quantify LPMO activity during protein purification. It is not suited to screen for LPMO activity in culture supernatants, to characterize mutational changes, or to measure physiological reactions. Despite its shortcomings, the Amplex red-based assay has been widely used and shows the need for a fast spectrophotometric assay to screen, produce, and purifiy LPMO.
Results and discussion
Screening for chromogenic substrates
Buffer and pH effects on LPMO activity
Cationic buffer based on pyridine, imidazole, and 2,4-dimethylimidazole result in LPMO activities as high as in the best anionic buffer (Fig. 3b). The pH profile of histidine, a zwitterionic species, is the only one which is monotonically decreasing between pH 5.5 and 7.0, indicating that the deprotonation of the imidazole ring inhibits LPMO activity, possibly by coordination of the active-site copper atom. Based on these results, it is concluded that many buffers can be used for the assay. We suggest the non-harmful sodium succinate/phosphate buffer with a high buffer capacity over a broad pH range of 4.5–8.0 (Fig. 3c).
As already indicated by the results with di- and tricarboxylic acids, the ionic strength of the buffer also influences the LPMO activity (Fig. 3d). The low ion concentration in 5 mM buffers lead to a twofold increase of activity over the 100 mM buffers either at pH 6.0 or 7.5. However, further experiments were performed with the 100 mM sodium succinate/phosphate buffer to ensure a high buffer capacity in the activity assay.
A problem observed in 2,6-DMP-based activity assays is the reaction of 2,6-DMP radicals with the final, colored product coerulignone, which decreases the absorbance and thereby affects the assay. This effect is also observed in laccase and peroxidase assays, but rarely covered in publications. This reaction cannot be stopped, but suppressed by using low 2,6-DMP concentrations and a low enzymatic activity in the assay. We suppose that a high concentration of the formed 2,6-DMP radical intermediates is responsible for the subsequent decolorization of coerulignone possibly by a polymerization reaction [20, 21, 22, 23]. On the other hand, higher 2,6-DMP concentrations increase the reaction rate of LPMO. Therefore, various 2,6-DMP concentrations at pH 6.0 and 7.5 were examined to find an appropriate range. The applicable 2,6-DMP concentration varies with pH and lies between 1–25 mM at pH 6.0 and 0.5–2 mM at pH 7.5. The 2,6-DMP concentration of 1 mM was used as a compromise to achieve reasonable enzymatic activity and assay stability. In cases where the assay should be as sensitive as possible, the 2,6-DMP concentration can be increased to the pH-dependent upper limit, but a shorter assay time has to be chosen to avoid the faster onset of the coerulignone decolorization reaction. When the assay is established for a new LPMO, it is important to evaluate this possible influence of the decolorization reaction. The assay is not affected when the increase of absorbance at 469 nm under steady-state conditions is strictly linear. A way to make the assay more sensitive would be the usage of hydrocoerulignone instead of 2,6-DMP, which, at a 1 mM substrate concentration has 15 times faster reaction rate (Fig. 2d). Unfortunately, hydrocoerulignone is difficult to solubilize in water. The faster hydrocoerulignone conversion also indicates that the 2,6-DMP oxidation is the rate limiting step in the assay and ensures that the assay measures the oxidation rate of 2,6-DMP.
Absorption coefficient and stoichiometry
The stoichiometry of the assayed LPMO reaction was determined in spectrophotometric cuvettes by following the conversion of substrate and cosubstrate in various concentrations. To this purpose, we assumed first the reaction stoichiometry to be similar to horseradish peroxidase, which is reported to be 1:1 for 2,6-DMP and H2O2. The same reaction stoichiometry for the LPMO-catalyzed conversion of 2,6-DMP and H2O2 has been confirmed by comparing the obtained results with horseradish peroxidase as catalyst (Fig. 4a). Because this LPMO reaction is similar to the peroxidase-catalyzed reaction, we call it peroxidase activity, until investigated in more detail. It was found that two 2,6-DMP molecules undergo a one-electron oxidation into the corresponding phenoxy radicals and donate in total two electrons to LPMO, which are transferred upon the acceptor hydrogen peroxide (Fig. 1). Up to a conversion of 20 µM 2,6-DMP, the measurements with LPMO and horseradish peroxidase (the reference reaction) correlate perfectly with the 2,6-DMP:H2O2 stoichiometry of 1:1 (Fig. 4a), but deviate from it at higher 2,6-DMP concentrations because of the coerulignone decolorization reaction described above. Figure 3b shows 2,6-DMP conversion by laccase at pH 6.0 and 7.5 (without H2O2), which also correlates well with the data in Fig. 3a. The two 2,6-DMP radicals dimerize into one hydrocoerulignone molecule, which is spectroscopically difficult to discriminate from 2,6-DMP. Hydrocoerulignone also reacts with H2O2 in an LPMO-catalyzed reaction (Fig. 4c, d) or with laccase in the absence of H2O2 (Fig. 4d) and forms the chromogenic product coerulignone with a stoichiometric ratio of 1:1.
Kinetics of the LPMO reaction
The peroxidase activity was found for three different LPMOs from two different organisms. The specific activity of purified enzyme preparations was measured at standard conditions: Neurospora crassa LPMO9C: 32.3 ± 0.9 U g−1, LPMO9F: 2.2 ± 0.2 U g−1, and Myriococcum thermophilum LPMO (gene identifier Myrth2p4_000359): 30.9 ± 0.7 U g−1. The specific activity between these homogeneously purified LPMOs differs, which indicates a different peroxidase activity of the enzymes, but it demonstrates that this activity is present in at least some LPMOs. Bissaro et al. have shown that LPMO activity with H2O2 is concentration dependent  and Kuusk et al. derived a KM value of CBP21 for H2O2 which was 2.8 ± 1.3 µM . The different measured specific activities of the three LPMOs can therefore also indicate different affinities towards the cosubstrate.
Kinetic constants of NcLPMO9C for H2O2 and 2,6-DMP
Kinetic constants of NcLPMO9C for H2O2
Kinetic constants of NcLPMO9C for 2,6-DMP
Cosubstrate 2,6-DMP (mM)
Vmax (U g−1)
Cosubstrate H2O2 (µM)
Vmax (U g−1)
34.3 ± 2.4
7.5 ± 0.2
245 ± 74
270 ± 20
39.8 ± 2.5
32.6 ± 0.8
100 ± 13
235 ± 15
41.1 ± 2.7
59.5 ± 1.4
144 ± 28
352 ± 43
7.3 ± 0.9
22.3 ± 0.6
172 ± 49
2500 ± 400
10.7 ± 1.1
32.5 ± 0.8
132 ± 32
2250 ± 250
11.8 ± 0.6
50.2 ± 1.1
124 ± 31
2300 ± 300
We also found no significant reduction of 2,6-DMP conversion when oxygen was removed from the LPMO assay. The observed specific LPMO activity in the presence of H2O2 and O2 was 32.3 ± 0.9 U g−1, whereas in the glove box under anaerobic conditions a specific activity of 30.6 ± 1.8 U g−1 was obtained.
Stability of the assay and control experiments
In accordance to Bissaro et al. , we found that the stability of LPMO is affected by H2O2. At the low H2O2 concentration used in the assay (100 µM), no loss of LPMO activity was observed during the assay. However, in titration experiments (Fig. 4a) the biggest achieved total turnover number of NcLPMO9C with H2O2 was 53. From our observations, we conclude that LPMO is deactivated by H2O2, especially under turnover conditions.
2,6-DMP conversion rates in control reactions at pH 7.5
NcLPMO9C + 2,6-DMP + H2O2
32.3 ± 0.9
NcLPMO9C (heat inactivated) + 2,6-DMP + H2O2
0.2 ± 0.1
NcLPMO9C + 2,6-DMP + H2O2 + 20 µM EDTA
0.4 ± 0.4
NcLPMO9C + 2,6-DMP (no H2O2 added)
0.6 ± 0.4
NcLPMO9C + H2O2 (no 2,6-DMP added)
0.1 ± 0.1
CuSO4 + 2,6-DMP + H2O2
0.6 ± 0.3
CuSO4 + 2,6-DMP (no H2O2 added)
0.5 ± 0.4
2,6-DMP + H2O2 (no NcLPMO9C added)
0.1 ± 0.1
Assay detection limit
The LOD was determined from 12 different LPMO concentrations, which were measured with the LPMO assay in four, fully randomized technical repeats. Assuming a Gaussian distribution, 95% represents the observed LOB values or LOD low-concentration sample values that exceed the defined LOB, respectively. The remaining 5% of blank values are false positive and only 5% of the low-concentration samples will produce values below the LOB. When using 1.645 × SD, no more than 5% of the values should be less than the LOB . The LOD is the lowest LPMO concentration that can be discriminated from the blank reading with a significance of 0.99 and was calculated to be 0.0125 µM (0.43 µg mL−1) of NcLPMO9C.
LPMO activity assay protocol
- Step 1:
Prepare a 116 mM sodium succinate/phosphate buffer (or another suitable buffer) by titrating the pH to 6.0 or (for enhanced assay sensitivity) to pH 7.5. The final concentration of the buffer in the cuvette will be diluted to 100 mM. Also prepare a 10 mM 2,6-DMP stock solution (for increased sensitivity at pH 6.0 prepare a 100 mM stock solution) in highly pure water and a 5 mM H2O2 stock solution in highly pure water. All solutions should be stored separately and used within 12 h.
- Step 2:
Take 1 mL of sample from the culture supernatant and centrifuge for 3 min at 6000×g to remove cells and other solids. Carefully remove 500 µL of the clear supernatant from the sediment and transfer into a clean vial. If taken from a clear solution, the sample needs no centrifugation and a smaller volume suffices. Store on ice until use.
- Step 3:
Transfer 860 μL of buffer, 100 μL of 2,6-DMP stock solution, and 20 μL of the H2O2 stock solution into a cuvette and mix the solutions well. Incubate the cuvette for 15 min at 30 °C before continuing with Step 4.
- Step 4:
Transfer the cuvette into a thermostated sample holder of the spectrophotometer and add 20 μL of properly diluted LPMO. If the LPMO activity is low, higher sample volumes can be used, but the buffer volume and ionic strength have to be adapted accordingly. Measure the increase of absorbance at 469 nm for 300 s and use the correct enzyme factor based on the sample volume, the enzyme dilution, and the molar absorption coefficient of coerulignone (ε469 = 53,200 M−1 cm−1) to calculate the peroxidase activity of LPMO.
The validated LPMO assay procedure is an easy method to quickly follow recombinant LPMO production and purification and is 50 times more sensitive than the previously published Amplex red assay. The 2,6-DMP assay has the potential to broadly be used to enhance the performance of LPMO production and to achieve higher yields and purer LPMO preparations after purification by allowing a more precise fractionation and pooling of the enzyme. The assay can also be used to determine non-catalytic enzyme properties such as thermal unfolding of substrate binding measurements where the peroxidase activity is only used to discriminate active from inactive LPMO molecules or to measure a fraction of LPMO in solution versus the LPMO fraction bound to a polymeric substrate.
Materials and enzymes
All chemicals were of the highest purity grade available and were purchased from Sigma-Aldrich unless stated otherwise. Methanol was purchased from Merck, pyrocatechol from Fluka, and hydrocoerulignone from MP Biomedicals. The lytic polysaccharide monooxygenases NcLPMO9C (Accession Number EAA36362.1) and NcLPMO9F (CAD70347.1) from Neurospora crassa and LPMO from Myriococcum thermophilum (Myrth2p4_000358 sequence deposited on http://www.fungalgenomics.ca) were recombinantly expressed in Pichia pastoris X-33 according to Kittl et al. . Production in a 5-L laboratory fermenter and column purification were also performed according to this publication. The purity of the enzyme was verified by SDS-PAGE. Laccase from Botrytis aclada was produced in a 5-L fermenter using Pichia pastoris as expression host and column chromatography according to Kittl et al. . Horseradish peroxidase was purchased from Sigma-Aldrich and used without further purification.
Measurement of LPMO concentration and hydrogen peroxide concentration
The LPMO concentration was determined in a 0.3-mm quartz cuvette from the absorption at 280 nm using molar absorption coefficients and molecular masses of NcLPMO9C ε280 = 46,910 M−1 cm−1, 34,300 g mol−1; NcLPMO9F ε280 = 51,130 M−1 cm−1, 23,200 g mol−1; and Myrth2p4_000358 ε280 = 44,140 M−1 cm−1, 22,515 g mol−1. The H2O2 concentration was determined in a 10-mm quartz cuvette from the absorption at 240 nm using a molar absorption coefficient of ε240 = 43.6 M−1 cm−1.
Screening of chromogenic substrates
NcLPMO9C was used to probe different phenolic compounds with and without H2O2 in the presence of molecular oxygen to determine if H2O2 increases the reaction rate. All experiments were performed in 100 mM succinate/phosphate buffer at pH 6.0. The phenolic compounds were dissolved in water (sinapic acid in methanol, final assay concentration of methanol 2%; hydrocoerulignone in DMSO, final assay concentration of DMSO 2%) and transferred into a cuvette containing 100 mM buffer, 2 µM LPMO, 100 µM H2O2, and 0.2 mM of the phenol (final concentrations). In reference experiments not containing H2O2, water was added instead of the H2O2 solution. Oxidation of the phenols in the presence or absence of H2O2 was followed for up to 5000 s in a diode-array spectrometer using quartz cuvettes.
pH optimum of LPMO activity
NcLPMO9C was used to measure enzymatic activity with final concentrations of 100 mM buffer, 100 µM H2O2, 25 mM DMP, and 0.10–3.75 µM NcLPMO9C (for histidine buffer a 30 µM NcLPMO9C was used to measure activity more accurately). For the whole range of a buffer, the same LPMO concentration was used, and the concentration was only adjusted between different buffers to remain in a measurable range of activity. The anionic buffers were adjusted with sodium hydroxide and the cationic buffers with hydrochloric acid. All measurement were performed at 30 °C and the change in absorbance at 469 nm was followed in a Perkin Elmer Lambda 35 UV/Vis spectrophotometer in triplicates. From pH 3.5 to 7.5, it was not necessary to measure blank reactions without NcLPMO9C, but starting from pH 8.0, a color formation by auto-oxidation of 2,6-DMP is observed. Therefore, it was necessary to measure blank reactions and subtract their slope from the corresponding measurement containing NcLPMO9C. The slope of the initial, steady-state rate was used to calculate the specific activity by using the molar absorption coefficient of coerulignone (ε469 = 53,200 M−1 cm−1) and the NcLPMO9C concentration. The effect of the buffer concentration was measured with the buffer systems sodium succinate/phosphate and pyridine/imidazole chloride at concentrations between 5 and 100 mM.
To probe the described LPMO reaction, we tested the effect of EDTA, heat treatment, the absence of H2O2, 2,6-DMP, LPMO, and the presence of copper(II) sulfate instead of LPMO. LPMO was added to the solution with final concentrations of 1 mM DMP and 100 µM H2O2 and 0.5 µM LPMO at 30 °C and measured for 300 s in a diode-array spectrometer using plastic cuvettes. The reaction was followed at 469 nm and the slope and specific activity was calculated. Blank reactions were performed, but omitting LPMO or 2,6-DMP or H2O2. To denature LPMO by heat, LPMO was added to a final concentration of 0.52 µM to the buffer and incubated for 1 h at 100 °C. To remove the copper atom from the active site of LPMO, we used 1 mM (final concentration) EDTA and incubated 25 µM LPMO for 30 min at 25 °C. Afterwards, the loss of activity was verified by an activity assay. Reactions with 0.5 µM CuSO4 were performed at standard conditions, but without LPMO.
LPMO activity assay
The suggested standard conditions for the LPMO activity assay are a temperature of 30 °C, an H2O2 concentration of 100 µM, a 2,6-DMP concentration of 1.0 mM, a 100 mM sodium succinate/phosphate buffer at pH 7.5 for maximum robustness and sensitivity or a 2,6-DMP concentration of 10.0 mM at pH 6.0 to account more physiological conditions for some LPMOs, and a reaction time of 300 s. One unit of LPMO activity is defined as the conversion of 2 µmol 2,6-DMP or the formation of 1 µmol coerulignone (ε469 = 53,200 M−1 cm−1) per min under reaction conditions.
LPMO activity assay under anaerobic conditions
Experiments in the absence of oxygen were carried out in a glove box (Whitley DG250, Don Whitley Scientific, Shipley, UK) which was continuously flushed with a nitrogen/hydrogen mixture (90/10). Residual oxygen was removed with a built-in palladium catalyst and the generated water vapor was absorbed on silica gel. Absorption was recorded with an Agilent 8453 UV–Vis spectrophotometer equipped with a photodiode array detector. All chemicals and reagents used within the glove box were extensively degassed by applying alternate cycles of vacuum and nitrogen pressure. Experiments were carried out in stirred cuvettes. Otherwise, the experimental setup was identical to the measurements carried out at ambient conditions. Experiments were carried out in three independent repeats.
Kinetic constants of LPMO and temperature effects
Kinetic constants were determined in 100 mM succinate/phosphoric acid buffer at pH 6.0 and 7.5. All experiments were performed in triplicates. The resulting curves were fitted to the Michaelis–Menten equation by non-linear least-square regression using SigmaPlot 12.5 (Systat Software, Chicago, Illinois, USA). For measurements performed at pH 6.0 2 µM, LPMO was used and for measurements at pH 7.5, 0.125 µM LPMO was used. To test the effect of the reaction temperature, the standard assay was preincubated at temperatures between 25 and 60 °C and 0.5 µM LPMO was added to start the reaction in the photometer.
Limit of detection
To determine the lowest concentration of LPMO which can be measured, we determined the limit of detection (LOD) for the standard LPMO assay according to . Twenty-four blank reactions without LPMO were measured to calculate the limit of blank (LOB). For the LOD measurement, four independent LPMO dilution series were prepared and measured in a completely randomized measurement scheme, which was prepared using the RAND function in Microsoft Excel 2016 (Microsoft Cooperation, Redmond, WA, USA). Quadruplets were measured to calculate the specific activity. A 6-time higher volumetric activity compared to the standard error of the blank was used to determine the LOD. To recalculate the LPMO concentration, the average activity of all measurements in the linear range was used.
Reactant stoichiometry and molar absorption coefficients of 2,6-DMP and hydrocoerulignone
NcLPMO9C, laccase, and horseradish peroxidase activity on 2,6-DMP was compared. The maximally formed absorption at 469 nm of reaction products at pH 6.0 and 7.5 in 10 mM sodium succinate/phosphate buffer was determined for all three enzymes and the molar absorption coefficient for 2,6-DMP and hydrocoerulignone was calculated. Final concentrations for of 2.5–50 µM H2O2 with 1.0 mM 2,6-DMP or 200 µM hydrocoerulignone and 2.5–50 µM 2,6-DMP or 1.25–10 µM hydrocoerulignone with 100 µM H2O2 were used and converted with 0.38 µM NcLPMO9C, 0.2 µM laccase, and 0.02 µM HRP. In case of laccase no H2O2 was added. The molar absorption coefficient of coerulignone was calculated from experiments of all three enzymes considering only substrate concentrations in the linear range.
Measurements were performed at 30 °C using a 4-mL electrochemical cell. The system setup involved a rotating disk electrode made from glassy carbon working electrode, a Ag|AgCl (3 M) reference electrode, a platinum counter electrode, an Autolab Rotator (RDE80739), an Autolab controller, and an Autolab potentiostat (PBSTAT204). The system was controlled using the NOVA 1.11 program from Autolab. Before measurements, the glassy carbon disk electrode was polished with an aluminum oxide suspension (Buehler; Master Prep Polishing Suspension, 0.05 μm). The rotating disk electrode was set to 250 rpm before starting the cyclic voltammetry measurement between 0 and 600 mV vs. the reference electrode with scan rate of 3 mV s−1. For the measurement of individual 2,6-DMP concentrations, 5 mL of a 100 mM sodium phosphate buffer, pH 6.0, was added to the cell. The buffer was left to equilibrate the system for 5 min. Different concentrations (0.03, 0.1, 0.3, 1.0, 5.0, and 20.0 mM) of 2,6-DMP were added to the cell, and after mixing for 5 min the measurements were started. The same procedure was performed for measurements at pH 8.0 using 100 mM sodium phosphate buffer, pH 8.0.
EB and RL developed the enzyme assay. MH performed kinetic measurements and assay validation. AH performed electrochemical measurements. CL and RK analyzed experimental data. EB coordinated the experiments and drafted the manuscript. BS, CO, and RL initiated, designed, and coordinated the study and wrote the final version of the manuscript. All authors read and approved the final manuscript.
This work was supported by the Austrian Science Fund project (FWF I2385-N28), the doctoral program “BioTop—Biomolecular Technology of Proteins” (FWF W1224), the Erasmus+ program which provided a scholarship to MH, and the WTZ project HR 12/2016 from ÖAD.
The authors declare that they have no competing interests.
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