A colorimetric assay to rapidly determine the activities of lytic polysaccharide monooxygenases
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Lytic polysaccharide monooxygenase (LPMOs) are enzymes that catalyze the breakdown of polysaccharides in biomass and have excellent potential for biorefinery applications. However, their activities are relatively low, and methods to measure these activities are costly, tedious or often reflect only an apparent activity to the polysaccharide substrates. Here, we describe a new method we have developed that is simple to use to determine the activities of type-1 (C1-oxidizing) LPMOs. The method is based on quantifying the ionic binding of cations to carboxyl groups formed by the action of type-1 LPMOs on polysaccharides. It allows comparisons to be made of activities under different conditions.
Based on the colorimetric detection and quantification of the pyrocatechol violet (PV)-Ni2+ complex, we have developed an assay to reliably detect and quantify carboxylate moieties introduced by type-1 LPMOs. Conditions were optimized for determining the activities of specific LPMOs. Comparisons were made of the activities against cellulose and chitin of a novel AA10 LPMO and a recently reported family AA11 LPMO. Activities of both LPMOs were boosted by hydrogen peroxide in the 1st hour of the reaction, with a 16-fold increase for the family AA11 LPMO, and up to a 34-fold increase for the family AA10 LPMO.
We developed a versatile colorimetric cation-based assay to determine the activities of type-1 LPMOs. The assay is quick, low cost and could be adapted for use in industrial biorefineries.
KeywordsLytic polysaccharide monooxygenase Enzyme assay Biomass deconstruction
LPMOs were first reported in 2010 . Despite intensive research, measurement of their activities remains a problematic task largely hampered by the low solubility of the products and because cleaved oligosaccharides can be re-absorbed back on to the surface of insoluble polysaccharides. Indeed, conventional reducing sugar assays are unsuitable for the detection of the products. Although techniques such as X-ray photoelectron spectroscopy (XPS) can detect C(=O)OH functional groups on insoluble polysaccharides introduced by type-1 LPMO, it has a low throughput and limited quantitative precision. Other currently available assay methods for LPMOs require laborious procedures such as a post-treatment with polysaccharide hydrolases to convert the insoluble oxidized polysaccharides into small, soluble oligosaccharides each with a reducing-end aldonic acid. These oligosaccharides are then separated and quantified by high-performance anion-exchange chromatography with pulsed-amperometric detection (HPAEC-PAD) using a strong alkali eluent [7, 8]. Such methods are accurate but time-consuming and require a library of aldonic acid oligosaccharide standards derived from different polysaccharides, limiting the study of LPMO activity to specialized laboratories. Several other LPMO assays have also been reported, including one in which chitin was radiolabelled with 14C on C-2 and the quantity of soluble 14C oligosaccharides released by LPMO activity determined . A microplate-based assay has also been developed that is based on labelling of reducing-end aldonic acid with a fluorescence dye . The chromogenic substrate 2,6-dimethoxyphenol (2,6-DMP) has also recently been used to quantify H2O2 consumption by LMPOs. The LPMO catalyzes the oxidation of 2,6-DMP to form the chromogenic product coerulignone at the expense of two H2O2. However, the result did not correlate with the oxidative activity on the polysaccharide substrates . A more convenient method of screening and comparing LPMO oxidative activities on different polysaccharide is urgently needed. Because oxidation at C1 by type-1 LPMOs generates aldonic acid at the reducing ends (Scheme 1), this contributes to the overall negative charge on the treated polysaccharide surface. This led us to develop an ion adsorption/desorption assay to quantify type-1 LPMO activities.
Results and discussion
Optimization of the stability of the Ni2+-pyrocatechol violet (PV) complex
Detection of LPMO AA11 activity using the cation-based assay
Detection of the activity of a novel LPMO AA10 using the cation-based assay
For comparing the concentrations of carboxylate moieties introduced by type-1 LPMOs under different conditions, this Ni2+ cation-based assay was found to be rapid and reliable. We demonstrated the Ni2+ cation-based assay is useful for measuring LPMO activity; multiple measurements can be achieved in 1 h on a 96-well plate. Compared with other reported assays, no post-enzymatic treatment, radioactive tracer or chemical labelling is needed. The cation-based assay has the potential to become a general method for comparing the activities of proteins within other LPMO families, such as the starch-specific AA13 family  or the xylan-specific AA14 family , or for investigating LPMO proteins within families with polysaccharide substrate specificity that are waiting to be discovered.
There are also limitations to this Ni2+ cation-based assay. First, we observed a discrepancy in the stoichiometric amounts of reductant and product that is most likely due to the CM cellulose standards we used for the carboxylate/Ni2+ calibration curve, or to the presence of other reductants from unexpected sources. Second, the Ni2+-carboxylate complex does not follow a 1:2 ratio as the spacing between the carboxylate moieties introduced by LPMO cannot be precisely regulated. For this reason, a better standard to simulate the aldonic acid introduced by LPMOs would certainly increase the reliability of the method. Finally, as this Ni2+ cation-based method does not detect C4-oxidized products generated by type-2 LPMOs, but it offers a way of distinguishing between LPMOs with type-1 and 2 activities.
In summary, a simple and stable colorimetric assay has been developed to investigate type-1 LPMO activity. This easily adaptable and scalable assay has the potential to be developed into a fully automated screening assay for LPMO oxidative activities against different polysaccharide substrates, including cellulose and chitin, and also water-soluble polysaccharides that can be precipitated by 95% ethanol, such as heteroxylans and xyloglucans. Finally, the assay can be scaled up for different applications in both academic and industrial settings that involve the synergistic use of polysaccharide hydrolases and LPMOs, which would allow their used to be further exploited and optimized.
Colorimetric measurements on the Ni2+–PV complex in different buffer solutions
NiCl2 solutions in 10 mM HEPES buffer (pH 8) or in ethanol:HEPES buffer (95:5 v/v) at 0–100 µM concentrations (500 µL each) were prepared. Each concentration was mixed with an equal volume of 400 µM pyrocatechol violet solution (PV, 500 µL) freshly made in 10 mM HEPES buffer or in ethanol:HEPES before use. The solutions were vigorously mixed and their absorbance at 650 nm was recorded immediately and every 5 min for 80 min in total. Correlation curves and equations were obtained by linear fitting. Other buffers were also tested, but we found the ethanol:HEPES buffer (95:5 v/v) provided the most stable Ni2+-carboxylate complex.
Colorimetric measurement of the reduction in concentration of the Ni2+ cation caused by ionic adsorption to carboxyl groups
In an optimized procedure, CM-cellulose (or AA10 and AA11 treated polysaccharides) was added to 540 µL ethanol-HEPES buffer (ethanol:10 mM HEPES, pH 8 = 95:5, v/v) giving CM-cellulose concentrations of 1.05, 2.71, 3.89, 5.05 and 7.03 mM. The solutions were mixed well and left undisturbed for 2 min. NiCl2 in ethanol:HEPES buffer (2 mM, 60 µL) was then added to the CM-cellulose solutions. After vigorous mixing, the solutions were left at ambient temperature for 2 min, and then centrifuged (16,000×g, 5 min) to precipitate Ni2+–CM cellulose particles. The supernatant (500 µL) was removed and mixed vigorously with PV (500 µL, 80 µM), giving a final PV concentration of 40 µM. The absorbance was recorded immediately. A standard curve was plotted from CM cellulose with a known degree of substitution (DS) using absorbance and concentration of carboxyl group (in the initial 600 µL = 540 µL of buffer + 60 µL of 2 mM NiCl2) derived from the above equation. Non-specific absorption of Ni2+ to Avicel (Sigma, MO, USA) and α-chitin (Sigma, MO, USA) was also examined using the procedure described above.
Preparation of FfAA11 and novel CmLPMOAA10
Primers used for cloning of CmAA10
pelB leader forward
The purified LPMO was saturated with copper by incubation with a threefold molar excess of CuCl2 for 1 h at 30 °C. Excess copper was removed by desalting the protein using a PD MidiTrap G-25 desalting column (GE Healthcare). The reactions were set up with 1% (w/v) cellulose substrates including the crystalline cellulose preparation Avicel (Sigma, MO, USA), and the amorphous cellulose PASC prepared as described by Zhang et al. . The substrates were incubated in Cu2+ saturated CmAA10 in 1 mM ascorbate, and 50 mM sodium phosphate buffer (pH 6.0). After 16 h incubation at 37 °C, the mixture was centrifuged and the supernatant collected for further analysis.
Analyses of enzymatic reaction products were performed by MALDI‒TOF MS (Applied Biosystems, CA, USA). The reaction product (5 µL) was mixed with 10 mM NaCl (3 µL) and 2,5-dihydroxybenzoic acid (10 mg/mL, 5 µL) in 50% (v/v) acetonitrile . Then 1 µL of the mixture was spotted onto a stainless steel plate and rapidly dried under vacuum for homogeneous crystallization. The spectrometry was done using an accelerating voltage of 20,000 V with a delay time of 200 ns. The spectrometer was operated in the reflection mode.
Study of the effect of H2O2 on AA10 and AA11 activities using the Ni2+ cation ionic adsorption assay
To study the effect of H2O2, multiple reactions were set up with 1% (w/v) α-chitin as substrate and 2 µM of FfAA11 (in 50 mM ammonium acetate, pH 5.0, 1 mM ascorbic acid), with H2O2 (0, 50, 100, 200 µM). The oxidized α-chitin was centrifuged (4000×g, 5 min), and the precipitate washed 6 times with 1 mL HEPES buffer. The washes were to remove any soluble enzymes and Cu2+ ions that could affect Ni2+ binding. The soluble aldonic acids were also removed in the washing steps, and only the carboxylate moieties left behind on the insoluble recalcitrant polysaccharides were analyzed.
The amounts of carboxylate moieties introduced were then quantified by the colorimetric assay described above. For CmAA10 and Avicel, the reactions were set up with 1 µM enzyme, followed by the same assay for FfAA11 as described above. Controls were set up by incubating the substrate with copper-free enzyme (without the pre-incubation of the enzyme with the threefold molar excess of CuCl2), and without ascorbic acid and H2O2. The maximum reaction velocities (Vmax) and turnover numbers (TN) of the LPMOs were calculated from nonlinear regression plots using Origin 9.0 software (OriginLab, Northampton, MA, USA).
DW and YH conceived and designed the project. DW and JL conducted the experiments. DW, FA and YH analyzed the data. DW, AW, FA, and YH wrote the manuscript. All authors read and approved the final manuscript.
We would like to thank the Knut and Alice Wallenberg Foundation for financial support and Professor Philip J. Harris for critically reading the manuscript.
The authors declare that they have no competing interests.
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This work was supported by the grant from Knut and Alice Wallenberg Foundation.
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