A Lytic Polysaccharide Monooxygenase from a White-Rot Fungus Drives the Degradation of Lignin by a Versatile Peroxidase

The enzymatic biodegradation of structural polysaccharides is affected by the degree of delignification of lignocellulose during the white-rot fungal decay process. The lignin matrix decreases accessibility to the substrates for LPMOs. H₂O₂ has been studied as a cosubstrate for LPMOs, but the formation and utilization of H₂O₂ in the reactions still represent an intriguing focus of current research. Lignin-degrading peroxidases and LPMOs usually coexist during fungal decay, and therefore, the relationship between H₂O₂-dependent lignin-degrading peroxidases and LPMOs should be considered during the wood decay process. The current study revealed that white-rot fungal LPMOs may be involved in the degradation of lignin through driving a versatile form of peroxidase activity in vitro and that H₂O₂ generated by PoLPMO9A was preferentially used for lignin oxidation by lignin-degrading peroxidase (PsVP). These findings reveal a potential relationship between LPMOs and lignin degradation, which will be of great significance for further understanding the contribution of LPMOs to the white-rot fungal decay process.

ABSTRACT

Lytic polysaccharide monooxygenases (LPMOs), a class of copper-dependent enzymes, play a crucial role in boosting the enzymatic decomposition of polysaccharides. Here, we reveal that LPMOs might be associated with a lignin degradation pathway. An LPMO from white-rot fungus Pleurotus ostreatus, LPMO9A (PoLPMO9A), was shown to be able to efficiently drive the activity of class II lignin-degrading peroxidases in vitro through H₂O₂ production regardless of the presence or absence of a cellulose substrate. An LPMO-driven peroxidase reaction can degrade β-O-4 and 5-5′ types of lignin dimer with 46.5% and 37.7% degradation, respectively, as well as alter the structure of natural lignin and kraft lignin. H₂O₂ generated by PoLPMO9A was preferentially utilized for the peroxidase from Physisporinus sp. strain P18 (PsVP) reaction rather than cellulose oxidation, indicating that white-rot fungi may have a strategy for preferential degradation of resistant lignin. This discovery shows that LPMOs may be involved in lignin oxidation as auxiliary enzymes of lignin-degrading peroxidases during the white-rot fungal decay process.

IMPORTANCE The enzymatic biodegradation of structural polysaccharides is affected by the degree of delignification of lignocellulose during the white-rot fungal decay process. The lignin matrix decreases accessibility to the substrates for LPMOs. H₂O₂ has been studied as a cosubstrate for LPMOs, but the formation and utilization of H₂O₂ in the reactions still represent an intriguing focus of current research. Lignin-degrading peroxidases and LPMOs usually coexist during fungal decay, and therefore, the relationship between H₂O₂-dependent lignin-degrading peroxidases and LPMOs should be considered during the wood decay process. The current study revealed that white-rot fungal LPMOs may be involved in the degradation of lignin through driving a versatile form of peroxidase activity in vitro and that H₂O₂ generated by PoLPMO9A was preferentially used for lignin oxidation by lignin-degrading peroxidase (PsVP). These findings reveal a potential relationship between LPMOs and lignin degradation, which will be of great significance for further understanding the contribution of LPMOs to the white-rot fungal decay process.

INTRODUCTION

Fungal deconstruction of lignocellulose is critical to carbon cycling in terrestrial ecosystems (1–3). White-rot fungi have been considered to be the most efficient wood decomposers in nature. They can depolymerize rigid lignin covalently associated with polysaccharides in plant biomass utilizing lignin-degrading class II peroxidases manganese peroxidase (MnP), versatile peroxidase (VP), and lignin peroxidase (LiP) (4), and then the exposed cellulose can be further synergistically hydrolyzed by a variety of extracellular glycoside hydrolases (5). There has recently been great interest in the lytic polysaccharide monooxygenases (LPMOs) that are widely distributed in white-rot fungi (6, 7). These copper-dependent enzymes can break glycosidic linkages on the cellulose surface through the hydroxylation of C₁ or C₄ carbon in glycosidic bonds, thus boosting the activity of glycoside hydrolase (7–10). The reaction of LPMOs was proposed to require the reduction of active-site copper with oxygen or hydrogen peroxide as the cosubstrate and to involve hydrogen atom abstraction by Cu-superoxide or Cu-oxyl intermediates, but the details of the catalytic mechanism are still unclear (11, 12).

The formation and utilization of H₂O₂ in LPMO reactions represent an intriguing focus of the current research. In the absence of polysaccharide substrates and in the presence of electron donors, the reduced active-site copper of non-substrate-bound LPMOs is known to lead to an uncoupled reduction of O₂, and then generate H₂O₂ (13–15). In the past, peroxide formation was thought to be a harmful side-reaction that results in LPMO self-inactivation due to the oxidative damage to residues near the copper site (12), but the same recent study reported that LPMOs could use H₂O₂ as a cosubstrate to oxidize polysaccharides (12, 15). It is obvious that the reactive oxygen species released from the Cu-oxygen intermediate of unbound LPMOs would be the source for peroxygenase reactions of LPMOs that are bound to substrates (15, 16). However, H₂O₂ formation could exceed its consumption due to weak binding between LPMOs and polysaccharide substrates (17). More importantly, the resistant lignin matrix surrounding cellulosic microfibrils could hamper the binding of LPMOs during white-rot fungal decay of lignocellulosic biomass (18–20), generating additional H₂O₂. White-rot fungi contain abundant diverse genes encoding LPMOs that are often extracellularly expressed during lignocellulose decay (21), and fungal decay leads to the production of abundant electron donors for LPMOs reduction, including lignin-derived compounds and other small molecule reductants such as hydroquinone and cellobiose dehydrogenase (CDH) (22). The reduction of considerable amounts of unbound LPMOs may generate excess H₂O₂ that damages LPMOs, which indicates that LPMOs could always suffer from self-inactivation during fungal decay if the H₂O₂ generated by LPMOs cannot be shunted.

It is well known that lignin-degrading peroxidases use H₂O₂ as a cosubstrate to mineralize the lignin. Multiple extracellular enzymes, such as glyoxal oxidases (GLOX), aryl-alcohol oxidases (AAO), and copper radical oxidases (23–25), generate H₂O₂ for lignin degradation. Some studies showed that multiple copies of LPMOs are coexpressed with peroxidases during white-rot fungal decay (21). Therefore, the issue arises as to whether LPMOs can drive the lignin degradation process of class II peroxidases through the presence of H₂O₂ generated by non-substrate-bound enzymes. Because LPMOs produce H₂O₂ at a relatively low rate in comparison with other well-known H₂O₂-generating enzymes (15), whether unbound LPMOs are involved in the reaction of lignin-degrading peroxidases with lignin remains to be determined in the absence of polysaccharide substrates. Moreover, it is unclear whether lignin degradation with lignin-degrading peroxidases is still driven by LPMOs in the presence of polysaccharide substrate. Clarifying the relationship between LPMOs and lignin degradation will be of great significance for further understanding the contribution of LPMOs to the decay process.

In this study, PoLPMO9A, the LPMO9A gene from Pleurotus ostreatus, a well-known white-rot fungus with great lignin-degrading ability, was expressed and the recombinant protein was produced. The results demonstrated that PoLPMO9A could drive the catalytic reaction of the peroxidase from Physisporinus sp. strain P18 (PsVP) (class II lignin-degrading versatile peroxidase) in vitro, regardless of the presence or absence of a cellulose substrate. Moreover, in the presence of PsVP, H₂O₂ generated by PoLPMO9A was preferentially utilized for lignin oxidation rather than for cellulose oxidation. The potential contribution of LPMOs involved in lignin degradation will be of great significance for further understanding the white-rot fungal decay process.

RESULTS

Expression of the recombinant PoLPMO9A and H₂O₂ production.

Pleurotus ostreatus LPMO gene PoLPMO9A was successfully inserted into the genome of Pichia pastoris X33 by electroporation and expressed using methanol as an inducer. PoLPMO9A protein production was confirmed in protein samples collected at different induction days by SDS-PAGE analysis (see Fig. S1A in the supplemental material). The molecular size of recombinant PoLPMO9A was approximately 25 kDa, which was consistent with the predicted molecular weight of the protein. The level of expression of PoLPMO9A gradually increased with induction time. The recombinant protein was further purified by anion-exchange chromatography (Fig. S1B). SDS-PAGE showed a single peptide after purification, which indicated that pure PoLPMO9A had been obtained. For qualitative analysis of the oxidative activity of PoLPMO9A, matrix-assisted laser desorption ionization–time of flight/time of flight mass spectrometry (MALDI-TOF/TOF MS) was performed with phosphoric acid-swollen cellulose (PASC) as a substrate. The results showed that PoLPMO9A was active on PASC, producing a series of cellulose oligosaccharides and corresponding oxidized oligosaccharides ranging from DP₅ox to DP₈ox (as shown in Fig. S2). The typical oxidized oligosaccharides were discovered and used to identify the oxidative activity of PoLPMO9A. Possible products in the heptamer ion clusters included the native Glc7 ([M+Na] = m/z 1,175.23), the C₁-oxidized lactone or C₄-oxidized ketoaldose (anhydrated species; [M+Na] = m/z 1,173.19), the C₁-oxidized aldonic acid or C₄-oxidized gemdiol (hydrated species; [M+Na] = m/z 1,191.23), the sodium adduct of the aldonic acid sodium salt ([M + 2Na-H] = m/z 1,213.37), the double oxidized products ([M+Na] = m/z 1,189.23), and the sodium adduct of the sodium salt of the double oxidized heptamer ([M + 2Na-H] = m/z 1,211.31) (26, 27). The data showed that PoLPMO9A yielded a C₁ oxidation product, because only this oxidation produces the sodium salt of aldonic acid (m/z 1,213.37 and 1,211.31), and that a C₄-oxidation product was also produced due to the strong signals for double oxidized products (m/z 1,189.23 and 1,211.31), which showed that the obtained PoLPMO9A could be a C₁/C₄ oxidizer.

In the absence of polysaccharide and in the presence of the reductant ascorbic acid (Asc), levels of H₂O₂ generated by PoLPMO9A were determined using the Amplex Red reagent/horseradish peroxidase assay (12, 13). As shown in Fig. 1, H₂O₂ formation was accompanied by the gradual consumption of reductant and enhanced with increasing initial concentration of reductant, but maintained at very low levels (less than 5 μM) in the absence of PoLPMO9A. This indicated that H₂O₂ was predominantly generated by reduced LPMOs rather than the direct reaction of reductant with O₂. H₂O₂ yields reached the maximum level after 6 or 8 h of reaction time and then slightly decreased, which could have resulted from H₂O₂ consumption of the self-inactivation reaction mixture of LPMOs in the absence of substrate. The calculated maximum yields of H₂O₂ from reaction mixtures initially containing 250, 500, and 1,000 μM reductant were 83.7 ± 4.0, 147.1 ± 2.5, and 185.5 ± 1.0 μM, respectively. Production of H₂O₂ increased with increasing amounts of ascorbic acid. The average rates of H₂O₂ production (over 6h) were 0.21 ± 0.01, 0.41 ± 0.01, and 0.52 ± 0.00 μM/min in the presence of 250, 500, and 1,000 μM ascorbic acid, respectively, which showed that the levels were about 2.0-fold and 2.5-fold higher in the presence of 500 and 1,000 μM ascorbic acid than in the presence of 250 μM ascorbic acid.

FIG 1.

Time courses for H₂O₂ production and degradation of Asc. (A) Time courses for H₂O₂ production in the absence and presence of PoLPMO9A (4 μM). (B) Time courses for degradation of Asc in the absence and presence of PoLPMO9A (4 μM). All reactions were carried out with Asc (250 μM, 500 μM, and 1,000 μM) in malonate buffer (100 mM; pH 4.5) at 30°C. Asc, ascorbic acid.

Lignin-degrading peroxidase activity driven by LPMOs.

To clarify whether the oxidative reaction of class II lignin-degrading peroxidases could be driven by reduced LPMOs, lignin-derived aromatic compounds were selected to evaluate the activity of PsVP, a recombinant lignin-degrading peroxidase, driven by PoLPMO9A. The selected aromatic compounds, including p-coumaric acid, ferulic acid, and sinapic acid, were derived from p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) type lignin, respectively (28). The data in Fig. 2 (see also Fig. S3 and S4) show that PoLPMO9A or PsVP alone could not oxidize the lignin-derived compounds but that the aromatic compounds could be greatly degraded when PsVP was incubated with PoLPMO9A, just as seen with incubation of PsVP with the initial concentration (150 μM) of exogenous hydrogen peroxide.The degree of degradation of aromatic compounds by PoLPMO9A-driven PsVP increased with time. In the absence of polysaccharide substrate, more than 80% of p-coumaric acid, ferulic acid, and sinapic acid was degraded by the PoLPMO9A-driven PsVP reaction. Additionally, the average degradation rate of p-coumaric acid, ferulic acid, and sinapic acid reached 0.19 μM/min, 0.13 μM/min, and 0.18 μM/min, respectively. The degree of degradation of lignin-derived aromatic compounds increased with increasing concentrations of PsVP, and all lignin-derived aromatic compounds were entirely degraded when the PsVP concentration reached 300 U/liter (Fig. S5), indicating that LPMOs could provide sufficient H₂O₂ for the oxidative reaction of lignin-degrading peroxidases.

FIG 2.

Degradation of lignin-derived aromatic compounds by PoLPMO9A-driven PsVP in the absence and presence of Avicel. The degree of degradation of different structures (H, G, and S) of phenolic lignin-related aromatic compounds was detected by high-performance liquid chromatography (HPLC). For degradation of lignin-derived aromatic compounds by PoLPMO9A-driven PsVP, the reactions with PsVP or PoLPMO9A alone were set as control experiments. Negative-control experiments contained lignin-derived aromatic compounds alone. Positive controls contained lignin-derived aromatic compounds, PsVP, and initial exogenous H₂O₂ (150 μM). All reactions were performed at 30°C in 100 mM malonate buffer (pH 4.5) with 500 μM substrate.

Unexpectedly, the lignin-degrading peroxidases continued to be activated by LPMOs even in the presence of polysaccharide substrate (Fig. 2; see also Fig. S5). The degree of degradation of lignin-derived aromatic compounds also increased with increasing concentrations of PsVP in the presence of Avicel cellulose, and there was no difference observed in terms of the final degree of degradation of lignin-derived compounds irrespective of whether cellulose was added during the reaction (P < 0.01). As shown in the LPMO adsorption experiments (Fig. S6), the amount of LPMOs bound to cellulose accounted for only a small portion of the total enzyme amount. The maximum proportion of bound LPMOs in the total protein was approximately 23.3%, which indicates that most of the LPMOs did not bind to the surface of cellulose. Obviously, the substrate-unbound LPMOs could provide H₂O₂ either for the peroxygenase reaction of LPMOs or for lignin-degrading peroxidases. Interestingly, MALDI-TOF/TOF MS analysis showed that PoLPMO9A oxidized Avicel cellulose and produced cellulose oligosaccharides and corresponding oxidized oligosaccharides when lignin-degrading peroxidases were not added but that the oligosaccharide products were not detected in the reaction involving the LPMO-driven peroxidases mentioned above (Fig. S7). These results suggested that H₂O₂ generated by unbound PoLPMO9A was prone to utilization for lignin biodegradation, rather than for cellulose oxidation, in the presence of lignin-degrading peroxidases (PsVP). Thus, regardless of the presence or absence of a cellulose substrate, the reduced PoLPMO9A can always efficiently drive the catalytic reaction of versatile peroxidase (PsVP).

Degradation of lignin model dimers.

It is well known that lignin is a highly branched and highly resistant phenylpropanoid polymer that is linked by aryl ether bonds and carbon-carbon bonds (29–31). To clarify that PoLPMO9A could drive versatile peroxidase (PsVP) to degrade dimeric lignin model compounds, β-O-4 and 5-5′ lignin model dimers were synthesized and used as substrates for PoLPMO9A-driven PsVP reactions. Two lignin model dimers were significantly degraded by PoLPMO9A-driven PsVP, with degrees of degradation of 46.5% ± 1.8% and 37.7% ± 1.3% for β-O-4 lignin and 5-5′ lignin model dimer, respectively (Fig. 3; see also Fig. S8 and Table S2 in the supplemental material). The liquid chromatography/mass spectrometry (LC/MS) spectra of the products revealed the oxidative cleavage of lignin model dimers and the resultant monomeric products (Fig. 3; see also Fig. S9 in the supplemental material). For the β-O-4 lignin model dimer, the main identified aromatic reaction products were compounds II (302.9 Da, M⁺ + H), III (319.0 Da, M⁺ + H), and IV (217.1 Da, M⁺ + Na; Fig. 3A), reflecting the oxidation of the phenolic aromatic side chains and breakage of the β-O-4 coupling linkage. The β-O-4 lignin model dimer (I) (343.0 Da, M⁺ + Na) was oxidized first into compounds II and III followed by the cleavage of β-O-4 linkage, resulting in lignin monomer compound IV. The results are consistent with research reported by Kong et al. (32) and Zeng et al. (33) indicating that the VP can break the β-O-4 linkage. For the 5-5′ lignin model dimer (I) (328.9 Da, M⁺ + Na), a 5-5′ coupling product (II) (305.1 Da, M⁺ + H) and monomeric product (III) (190.9 Da, M⁺ + Na) were also observed (Fig. 3B). These types of products were highly similar to typical and well-known oxidation products in 5-5′ lignin model compound oxidations by laccase (34). The alcohol group of the 5-5′ lignin model dimer was oxidized to aldehyde (II), and then compound II was depolymerized by breaking the 5-5′ coupling linkage, leading to production of monomer compound III.

FIG 3.

Degradation of the lignin model dimers by PoLPMO9A-driven PsVP and LC/MS spectra of the products. Panels A and B present the degree of degradation and mass spectra of coupling products of β-O-4 and the 5-5′ lignin model dimer, respectively. For degradation of lignin model dimers by PoLPMO9A-driven PsVP, the reactions with PoLPMO9A or PsVP alone were set as control experiments. Negative-control experiments contained model dimers alone. Positive controls contained lignin model dimers, PsVP, and initial exogenous H₂O₂ (150 μM). All reactions were performed at 30°C in 100 mM malonate buffer (pH 4.5) with 500 μM substrate. Samples were subjected to HPLC and LC/MS analysis. The control spectrum is for the substrate in buffered solution. Intens, the degree of degradation of lignin model dimer.

Degradation of kraft lignin and natural lignin.

Two types of complex lignin polymers, namely, natural lignin isolated from bamboo culms and kraft lignin obtained from pulping wastewater, were further used as substrates to evaluate the efficiency of lignin degradation driven by the PoLPMO9A-driven PsVP reaction. Usually, kraft lignin has a lower molecular weight and higher abundance of resistant structures, such as 5-5′, α-5, diphenylmethane, and stilbene units, than natural lignin (29, 35). Pyrolysis gas chromatography mass spectrometry (Py-GC/MS) analysis was performed to characterize alterations in the lignin structure after PoLPMO9A-driven PsVP treatment. Analytic pyrolysis of lignin yielded a series of phenolic products, which were used to characterize the chemical structural change of trace lignin samples after enzyme delignification. Approximately 50 to 70 phenolic compounds were identified in the pyrolysis products of treated and untreated samples (Fig. 4; see also Table S3 and S4). These phenolic compounds were composed of S-type, G-type, and H-type lignin derivatives, as well as the catechol derivatives (1,2-benzenediol) that are the demethylated products of lignin. The amounts of S-type, G-type, and H-type lignin derivatives of the kraft and natural lignin that changed after PoLPMO9A-driven PsVP treatment were similar to those seen after PsVP treatment with initial H₂O₂ added.

FIG 4.

Determination of the distribution of H, G, and S units by pyrolysis GC/MS of untreated and two-enzyme treated lignin. Panels A and B represent kraft lignin and natural lignin, respectively.

G-type lignin derivatives were predominantly (>80%) products of kraft lignin derived from the wastewater of softwood pulping. The treatment of PoLPMO9A-driven PsVP led to a great decrease in the relative abundances of some G-type derivatives in the pyrolysates of kraft lignin, especially for 2-methoxy-4-methylphenol (peak 16^G; reduced by 63.6%). The total abundance of G-type derivatives was decreased by 15.1% after enzymatic treatment, which indicated that the aromatic ring of kraft lignin can be disrupted by enzymatic treatment. In addition, the pyrolysates of natural lignin of bamboo culms contained not only G-type derivatives but also S-type and H-type derivatives, and the G/S/H derivative ratio calculated for the control lignin sample was 37/56/5. After PoLPMO9A-PsVP treatment, the total abundances of H-type and S-type derivatives were reduced by 26.8% and 8.2%, and some H-type and S-type derivatives such as 4-hydroxy-acetophenone (peak 37), 4-hydroxy-benzenepropanoic acid (peak 54) trans-sinapyl alcohol (peak 68), and 3,5-dimethoxy-4-hydroxycinnamaldehyde (peak 69) completely disappeared. Moreover, enzymatic treatment led to a slight decrease in the total abundance of G-type derivatives and the estimated G/S/H ratio of pyrolysates of treated lignin was 40/55/4. A great increase in the abundance of some G-type derivatives, such as 4-[(1E)-3-hydroxy-1-propenyl]-2-methoxyphenol (peak 57) and 2-methoxyphenol (peak 5). This increase would have arisen from demethylation of the S-type unit by the PoLPMO9A-driven PsVP reaction, which was consistent with previous findings in the research of white-rot fungal delignification (36–38). On the basis of the data presented above, it was concluded that kraft lignin and natural lignin were efficiently degraded, leading to monomeric compositional variation of lignin unit distribution as a result of the PoLPMO9A-driven PsVP treatment, which indicates that LPMOs can provide a driving force to delignification with lignin-degrading peroxidases.

DISCUSSION

LPMOs have recently been considered to be important for the biodegradation of structural polysaccharides during the white-rot fungal decay process (38, 39). The binding of LPMOs onto the surfaces of crystalline cellulose is a prerequisite for substrate oxidation. However, the resistant lignin matrix coating cellulose microfibers can decrease cellulose accessibility to enzymes in the process of fungal decay (38), which limits the cellulose oxidation by LPMOs. More importantly, the presence of non-substrate-bound LPMOs can result in the formation of H₂O₂, leading to self-inactivation. In this study, we provided, to our knowledge, the evidence that white-rot fungal LPMO drives the fungal class II lignin-degrading peroxidases for lignin model compound biodegradation, including cleavage of lignin interunit linkages, disruption of aromatic rings, and demethylation of lignin. Considering the findings from other studies, our data suggested that LPMOs may have a wider role in white-rot fungal decay.

First, white-rot fungi have versatile extracellular H₂O₂-generating enzymes that participate in the lignin degradation reaction of class II peroxidase, such as aryl-alcohol oxidase (AAO) and glyoxal oxidase (GLOX) (23). The expression of H₂O₂-generating enzymes depends on the fungal isolate, wood substrate species, and physiological factors, and different H₂O₂-generating enzymes play roles in different stages of fungal decay (40, 41). Some H₂O₂-generating enzymes, such as glyoxal oxidases, are lacking in some fungi, including Ceriporiopsis subvermispora, Fomitiporia mediterranea, and Heterobasidion annosum, but almost all white-rot fungi reported thus far have contained LPMOs (25). Thus, non-substrate-bound LPMOs may also be potential providers of hydrogen peroxide for class II peroxidases during fungal decay, like other fungal H₂O₂-generating enzymes. Furthermore, some H₂O₂-generating enzymes require specific electron donors with a high oxidation potential, such as aromatic compounds for aryl-alcohol oxidase. However, LPMOs can be reduced with a wide range of sources of electron donors, including ascorbic acid, cellobiose dehydrogenase (CDH), phenols from lignin degradation or plant extractives, light-activated photosynthetic pigments, and even light-activated TiO₂ (42, 43).

Second, many of genes encoding the lignin-degrading peroxidases and LPMOs are usually simultaneously expressed during fungal decay of wood (39). LPMOs that are widespread and present in many copies, even with significantly more isoenzymes than some cellulases, have been identified in the genome of most ascomycetous and basidiomycetous (white-rot and brown-rot) fungi (44). A recent study reported that when the white-rot fungus Obba rivulosa, a selective lignin decomposer, was cultivated on solid spruce wood, the relative transcript levels per putative CAZy enzyme activity were compared during the cultivation. MnPs were the most highly expressed, followed by LPMOs (21). LPMOs seem to play potential roles in fungal delignification with class II peroxidases. Taking into account the multiple copies of LPMO that are coexpressed with lignin-degrading peroxidase, fungal LPMOs may have versatile functions, including boosting the hydrolysis of structural polysaccharides and providing H₂O₂ to peroxidase for lignin degradation.

Third, some recent reports have revealed the connections between enzymatic delignification and cellulose degradation with LPMOs. LPMOs can use aromatic compounds derived from fungal degradation as a reductant for fueling cellulose oxidation (22, 45). Our studies showed another possible connection between the fungal degradation of lignin and structural polysaccharide: lignin degradation with peroxidase could be conversely fueled by LPMOs. In the current study, the selected lignin-derived compounds were not oxidized in the presence of LPMOs or reduced LPMOs alone, which indicated that these compounds could not be degraded as electron donors of LPMOs and that their degradation in LPMO-driven peroxidase reactions should be attributed to the peroxidase reaction driven by H₂O₂ released from unbound LPMOs (see Table S2 in the supplemental material). Interestingly, the lignin-derived compound p-coumaric acid was almost completely degraded in the LPMO-driven peroxidase reaction, but its degree of degradation was approximately 38.1% ± 1.2% in the exogenous H₂O₂-driven peroxidase reaction. A recent study showed that degradation products (diphenols) of p-coumaric acid with polyphenol oxidase could be a satisfactory reducing agent for LPMOs (46). Lignin-degrading peroxidases and polyphenol oxidase also have partially similar catalytic functions for oxidation of lignin-derived compounds, leading to the formation of hydroxylated phenolic compounds. It is possible that the reduced LPMO could activate the peroxidase activity to attack p-coumaric acid, producing hydroxylated products that further serve as a reductant of LPMOs and are simultaneously degraded by the peroxidase and LPMO.

Currently, whether LPMOs utilize O₂ or H₂O₂ as a physiological cosubstrate remains open to debate (47), but it is clear that LPMOs work faster when provided with H₂O₂ (12, 47) and that they prefer H₂O₂ when presented with both cosubstrates (12). If H₂O₂ is a cosubstrate of LPMOs during white-rot fungal decay, it should be generated by H₂O₂-generating enzymes or non-substrate-bound LPMOs for the peroxygenase reaction of LPMOs. The present results revealed an interesting phenomenon, consisting of lignin-degrading peroxidase activity that was always driven by reduced LPMOs in vitro, regardless of the presence or absence of a cellulose substrate. Some studies reported the apparent lack of H₂O₂ production when LPMOs were incubated with substrates (1). In fact, it was not due to hydrogen peroxide not being produced. It may, however, be that H₂O₂ was simply being consumed in productive LPMO reactions with the substrate. H₂O₂ production was previously shown to depend on whether LPMOs were completely bound to the polysaccharide substrate (15). H₂O₂ produced by unbound LPMOs is still consumed to carry out H₂O₂-driven catalysis (15). The results of the current study showed that free LPMOs always existed due to incomplete binding of LPMOs to Avicel cellulose (see Fig. S6 in the supplemental material). The issue naturally arises as to whether H₂O₂ generated by unbound LPMOs was used for delignification or cellulose oxidation, because both the lignin-degrading peroxidases and LPMOs are secreted and expressed during fungal decay. Our results showed that oxidized products of cellulose were not detected and that lignin-derived compounds were still greatly degraded when LPMOs coexisted with the lignin-degrading peroxidase and both cellulose and lignin-derived compounds were used as substrates (Fig. S5 and S7). On the other hand, the K_m values for PsVP and PoLPMO9A with respect to affinity to H₂O₂ were 8.6 ± 2.8 μM and 3,661.0 ± 601.5 μM, respectively, which indicated that the affinity of PsVP to H₂O₂ was stronger than that of PoLPMO9A. This indicated that H₂O₂ released by PoLPMO9A is preferentially utilized for the delignification of lignin-degrading peroxidase (PsVP) instead of cellulose oxidation of PoLPMO9A. Some recent studies also showed that the formation of oxidized products from cellulose can be inhibited by horseradish peroxidase, a plant peroxidase (12, 47). These phenomena indicate that the peroxidase may competitively inhibit the LPMO activity on cellulose and that cellulose degradation boosted by LPMOs may be regulated by lignin degradation with class II peroxidases during fungal decay.

Many reports have confirmed that the biodegradation of structural polysaccharides with extracellular glycoside hydrolases is affected by delignification during white-rot fungal decay (38). However, there have been few published studies concerning the relationship between the lignin biodegradation reaction with lignin-degrading peroxidase and the catalytic reaction of LPMOs. On the basis of the present findings, we propose a possible mechanism of interaction between fungal LPMOs and class II lignin-degrading peroxidase during lignocellulose decay (Fig. 5). On one hand, the physical impediment of lignin limits the accessibility of LPMOs to the cellulose substrate, leading to the excess generation of H₂O₂ by non-substrate-bound LPMOs. The accumulated H₂O₂ could be further utilized by lignin-degrading peroxidase for lignin degradation, reducing the physical barriers of lignin, enhancing the binding of LPMOs to cellulose, and accelerating the cellulose oxidation by LPMOs. On the other hand, when lignin-degrading peroxidase and lignin substrate are present, H₂O₂ generated by the reduced non-substrate-bound LPMOs may be preferentially utilized for lignin degradation rather than for cellulose oxidation, regardless of the presence or absence of a cellulose substrate corresponding to LPMOs. This indicates that white-rot fungus could have a strategy of preferential degradation of lignin when LPMOs and class II peroxidase are coexpressed during fungal decay, which would contribute to overcoming biomass recalcitrance to the decay. Moreover, lignin-degrading peroxidases may regulate and limit H₂O₂ levels when excess LPMOs are expressed or left unbound to the polysaccharide substrate, preventing LPMOs self-inactivation caused by peroxide accumulation.

FIG 5.

Schematic representation of the degradation of lignocellulose. The resistant lignin matrix surrounding cellulosic microfibrils can hamper the binding of LPMOs. During white-rot fungal decay of lignocellulosic biomass, abundant electron sources for LPMO reduction are produced, including lignin-derived compounds and other small-molecule reductants such as ascorbic acid, hydroquinone, and cellobiose dehydrogenase, which could lead to the reduction of considerable amounts of unbound LPMOs and generation of excess of H₂O₂ to damage LPMOs. When LPMOs are coexpressed with lignin-degrading peroxidases, LPMOs can drive the lignin degradation process of class II peroxidases by the H₂O₂ generated by non-substrate-bound enzymes.

Overall, our present study confirmed that the white-rot fungal PoLPMO9A can drive lignin biodegradation with class II lignin-degrading peroxidase (PsVP) in vitro regardless of the presence or absence of a cellulose substrate and showed that H₂O₂ generated by the PoLPMO9A was preferentially used for lignin oxidation by lignin-degrading peroxidase rather than for cellulose oxidation. These results provide a basis for understanding the relationship between LPMOs and class II lignin-degrading peroxidase. It is likely that LPMOs play the role of auxiliary enzymes in activating class II lignin-degrading peroxidase during white-rot fungal decay and not just as auxiliary enzymes of glycoside hydrolases. These findings are of great significance for further understanding the contribution of LPMOs to the white-rot fungal decay process.

MATERIALS AND METHODS

Chemicals.

Commercial kraft lignin (lot no. MKBV5831V) and lignin-derived aromatic compounds (p-coumaric acid, ferulic acid, and sinapic acid) were purchased from Sigma-Aldrich (USA). RNA extraction kits and a PrimeScript reverse transcriptase (RT) reagent kit with genomic DNA (gDNA) eraser were obtained from TaKaRa. A ClonExpress MultiS One Step cloning kit was purchased from Vazyme. Natural lignin was prepared from bamboo. β-O-4 (48, 49) and 5-5′ lignin model dimers (50) were synthesized and characterized in our lab. All other chemicals used were analytical grade. Mentioned above, the kraft lignin product (kraft lignin) is isolated from a commercial pulp mill using predominantly Norway spruce as raw material. “Kraft lignin” implies the process by which lignin is separated from the cellulose by use of either a hot acid (sulfite) or hot alkaline (sulfate) method. Bamboo is a type of woody grass, with structural characteristics distinct from those of wood or other grasses. The chemical composition of bamboo culm, which has higher lignin content than grasses, is similar to that of hardwood. The natural lignin (milled wood lignin) from Phyllostachys pubescens culms was obtained and purified according to the classical procedure (51, 52). Grass meal prepared from bamboo culms was extracted with ethanol/toluene 1:2 (vol/vol). Then around 30 g of the grass meal was finely ball-milled in a Retsch PM100 planetary ball mill for 72 h at 400 rpm using an agate jar and agate balls of 2 cm diameter. The ball-milled grass meal was then extracted with dioxane/water (96:4, vol/vol) for three times. The suspension was centrifuged and the supernatant evaporated under reduced pressure. The obtained sample was redissolved in acetic acid/water (9:1, vol/vol). The solution was then added into cold water, and the precipitate was separated by centrifugation. The precipitate was dried and dissolved in 1,2-dichloroethane/ethanol (2:1, vol/vol). The mixture was then centrifuged to eliminate the insoluble material. The resulting supernatant was precipitated into cold diethyl ether, centrifuged, and washed with diethyl ether and petroleum ether. The purified natural lignin was obtained after dried under a current of N₂. The yield of the natural lignin was 24.8% of the original bamboo lignin content.

Strains and culture conditions.

A LPMO-encoding gene was cloned from P. ostreatus HAUCC 162. The fungus was maintained on a potato dextrose agar (PDA) slant at 4°C and activated for 7 days on a fresh PDA slant. The mycelia collected for RNA extraction were produced by solid-state fermentation in 250-ml flasks with corn stalk (substrate/H₂O = 1:2.7) at 28°C for 15 days. The recombinant plasmid pET 28a-PsVP was constructed for expression of the versatile peroxidase (PsVP) (GenBank accession no. KY293299.1) from Physisporinus sp. P18 and expressed in Escherichia coli Rosetta (DE3). Escherichia coli DH5α was used to clone the PoLPMO9A-encoding gene and was grown overnight at 37°C on LB medium (5 g yeast extract, 10 g tryptone, and 10 g NaCl, containing 20 g agar per liter). Pichia pastoris strain X33 used for heterologous expression of the PoLPMO9A gene was grown at 28°C on YPD medium (10 g yeast extract, 20 g peptone, 20 g glucose, and 20 g agar per liter). YPDS medium (10 g yeast extract, 20 g peptone, 20 g glucose, 182.2 g sorbitol, and 20 g agar per liter) containing 100 μg/ml Zeocin was used for selection of the transformants. Positive recombinant strains were cultivated and induced in liquid BMGY medium (10 g yeast extract, 20 g peptone, 100 ml 13.4% [wt/vol] YNB, 100 ml 10% [vol/vol] glycerol, 100 ml 1 M [pH 6.0] potassium phosphate buffer, and 2 ml 0.02% [wt/vol] biotin per liter) with 0.5% (vol/vol) methanol according to the manual of a Pichia expression kit (Invitrogen).

Cloning and heterologous expression of the gene encoding PoLPMO9A.

The gene encoding PoLPMO9A (protein identifier [ID]: 1098582) was predicted in the sequenced genome of Pleurotus ostreatus PC15 v2.0 (http://genome.jgi.doe.gov/). Total RNA was extracted using RNA extraction kits (TaKaRa), and first-strand cDNA was then synthesized using a PrimeScript RT reagent kit (TaKaRa). The open reading frame of the PoLPMO9A was analyzed, and the region encoding the mature peptide of PoLPMO9A was obtained by PCR amplification from first-strand cDNA using primers (forward) AGAGAGGCTGAAGCTGAATTCCACGGTTACGTCCCCCAAATAAAAATAGG and (reverse) GAGATGAGTTTTTGTTCTAGATTAAAATGCCAAGCCAGGATAGACTGGAC (with EcoRI and XbaI sites underlined, respectively) and then cloned into vector pMD-18T (TaKaRa, Japan) and sequenced. For expression, the correct gene fragment was inserted into pPICzαA vector (Invitrogen, USA) digested with EcoRI and XbaI using a ClonExpress MultiS One Step cloning kit, resulting in recombinant plasmid pPICzαA-PoLPMO9A.

After resequencing, the confirmed pPICzαA-PoLPMO9A plasmid was linearized by the use of PmeI and then transformed into Pichia pastoris X33 competent cells by electroporation according to the Pichia expression vector manual (Invitrogen). Positive strains were screened on the basis of Zeocin resistance using YPDS media with 100 μg/ml Zeocin and tested further for expression of PoLPMO9A with BMGY media at 180 rpm and 28°C according to the instruction manual of an EasySelect Pichia expression kit (Invitrogen). To maintain induction, methanol at a final concentration of 0.5% (vol/vol) was added to the culture every 24 h.

Enzyme and protein assay.

To identify the expression of the recombinant enzyme PoLPMO9A, the culture media were centrifuged at 4°C and 8,000 rpm for 10 min, and the supernatants were subjected to SDS-PAGE analysis using the samples collected at approximately 5 to 9 days of culture. Protein concentrations were analyzed using the Bradford method with bovine serum albumin as a standard (53). SDS-PAGE was done according to Laemmli’s method (54). Proteins were visualized by staining with Coomassie brilliant blue R-250.

For purification of PoLPMO9A, the supernatants present at 9 days of culture of the recombinant strains were collected via centrifugation at 8,000 × g for 10 min and then treated with ammonium sulfate to reach a final saturation of 80% (wt/wt) at 4°C for 16 h in a one-step procedure. The precipitates were collected via centrifugation at 12,000 rpm for 20 min, redissolved in 20 mM Tris-HCl buffer (pH 7.5), and then subjected to dialysis with 20 mM Tris-HCl buffer (pH 7.5) until ammonium sulfate was removed. The solution obtained from the dialysis procedure was concentrated and loaded onto an anion-exchange column preequilibrated with 20 mM NaCl-Tris-HCl buffer (pH 7.5). The column was then washed to remove any unbound protein with the same buffer. Proteins were subjected to an elution gradient with a 1 liter of 20 mM Tris-HCl buffer containing 1 M NaCl (pH 7.5) at a 1 ml/min flow rate (55). The fractions were collected to characterize LPMO activity. To verify the activity of PoLPMO9A, phosphoric acid-swollen cellulose (PASC), prepared using Avicel PH-101 (Sigma-Aldrich), was chosen as the substrate (56). The 1-ml-reaction-mixture system used contained 2 mg/ml PASC–0.05 mg/ml PoLPMO9A–2 mM ascorbic acid–20 mM ammonium acetate buffer (pH 5.0). The reaction mixtures were treated overnight at 28°C. The supernatant was purified and then used for the mass spectrometry analysis. For MALDI-TOF/TOF MS analysis, 1 μl volumes of samples were mixed with 1 μl of 10 mg/ml 2,5-dihydroxybenzoic acid (DHB)–50% acetonitrile–10 mM sodium acetate on a OPTI-TOF 384 target plate and analyzed by the use of a MALDI-TOF/TOF 5800 instrument (AB Sciex, USA) (26, 57).

For the expression and purification of PsVP (Protein ID: ARA74332.1), recombinant Escherichia coli Rosetta (DE3) containing expression plasmid pET 28a-PsVP was grown in LB liquid media containing 50 μg/ml ampicillin and 34 μg/ml chloramphenicol and induced with 1 mM IPTG (isopropyl-β-d-thiogalactopyranoside) for 8 h at 37°C. The cells were centrifuged at 4°C and 10,000 rpm for 5 min and resuspended in lysis buffer (50 mM Tris-HCl [pH 8.0], 10 mM EDTA, 5 mM dithiothreitol [DTT]) at 1/10 of the original culture volume, and lysozyme was added at a final concentration of 2 mg/ml. After 1 h of incubation at 4°C, 4 U/ml DNase I was added and the cells were disrupted using the high-pressure cracker at 8e+7 Pa) for 15 min. The lysed cells were centrifuged at 4°C and 12,000 rpm for 30 min. The supernatants were discarded and the inclusion bodies containing the recombinant protein washed 2 to 3 times with 20 mM Tris-HCl containing 1 mM EDTA, 5 mM DTT, and 2 M urea (pH 8.0) and then resuspended in 50 mM Tris-HCl buffer containing 1 mM EDTA, 1 mM DTT, and 8 M urea (pH 8.0) at 1/100 the original culture volume. The solution was treated at room temperature for 1 h to fully solubilize the inclusion bodies and then centrifuged, and the final-volume supernatants were adjusted to yield 0.1 mg/ml protein with 50 mM Tris-HCl buffer (pH 9.5) containing 30 μM hemin, 5 mM CaCl₂, 0.7 mM glutathione disulfide (GSSG), and 0.1mM DTT to refold the inactive protein (58, 59).

The refolding solution was placed overnight at room temperature in the dark and extensively dialyzed against 20 mM sodium acetate buffer (pH 4.3) containing 1 mM CaCl₂. Insoluble protein was removed by centrifugation at 4°C and 12,000 rpm for 30 min, and the supernatants were further dialyzed against 20 mM NaH₂PO₄-NaOH buffer (pH 7.4) and concentrated with polyethylene glycol. Refolded preparations containing recombinant PsVP with 0.5 M NaCl were purified using a Ni²⁺ His tag column (GE) according to the manufacturer’s instructions. Samples were loaded into a 1 ml Ni²⁺ His tag column that had been preequilibrated with 50 mM NaH₂PO₄-NaOH buffer (pH 7.4) containing 0.5 M NaCl. The column was washed with 50 mM NaH₂PO₄-NaOH buffer (pH 7.4) containing 0.5 M NaCl and 5 mM imidazole to remove any unadsorbed protein. PsVP was eluted with 50 mM NaH₂PO₄-NaOH buffer (pH 7.4) containing 0.5 M NaCl and 50 mM imidazole. Fractions containing PsVP activity were pooled, dialyzed against 10 mM sodium tartrate (pH 5.0), and stored at −20°C (58, 59). The activity of PsVP was determined by monitoring the formation of Mn(III)-malonate complex at A₂₃₈ (ε₂₃₈ = 6,500 M⁻¹cm⁻¹). One activity unit was defined as the amount of enzyme oxidizing 1 μmol of substrate per min. The reaction mixture contained 1 mM MnSO₄ and PsVP and 100 mM sodium malonate (pH 4.5) with the addition of 0.1 mM H₂O₂ to start the reaction (60). SDS-PAGE showed a single peptide after purification using a Ni²⁺ His tag column as shown in Fig. S1C in the supplemental material, which indicated that pure PsVP had been obtained. After refolding and in vitro purification, the activity of PsVP reached 19.8 U/mg.

Assay for H₂O₂ production and ascorbic acid.

Ascorbic acid (Asc) was used as a reducing agent for PoLPMO9A to evaluate H₂O₂ production. To determine the effect of Asc on H₂O₂ production, the reactions were performed with 4 μM PoLPMO9A and Asc mixed with 100 mM malonate buffer (pH 4.5). Different concentrations of Asc (250 μM, 500 μM, and 1,000 μM) were selected to evaluate H₂O₂ production by PoLPMO9A. The H₂O₂ production was quantified by using the Amplex Red/horseradish peroxidase assay according to a published protocol reported by Bissaro et al. (12, 13). For each reaction mixture, 50 μl was sampled at regular intervals and mixed with 50 μl of NaOAc (50 mM) (pH 4.5). To determine the H₂O₂ concentration, 50 μl of the mixture (or dilutions of it) was mixed with 50 μl of a premix containing 50 μM Amplex Red (Invitrogen, USA) and 7.14 U/ml peroxidase. The reaction mixture (100 μl) was incubated in a 96-well plate for 10 min before recording the absorbance at 560 nm. The standard curve was prepared using H₂O₂ solutions. A UV spectrophotometric method was used for monitoring the concentration of Asc. Asc was quantified by converting the maximum absorbance value (at 265 nm) using an Asc standard curve. For all conditions, a reaction mixture lacking PoLPMO9A was prepared and subjected to the same reaction conditions and sample treatments. All the results represented in Fig. 1 are the means and standard deviations of data from three replicates (independent biological samples). Means and standard deviations were calculated using OriginPro 8.0.

Degradation of lignin by LPMO and class II lignin-degrading peroxidase.

Three monomeric lignin-derived aromatic compounds were used as substrates to evaluate the activity of LPMO-driven class II lignin-degrading peroxidase reactions in the absence and presence of Avicel cellulose. The reactions for single incubation were performed for 48 h with substrates (500 μM) with or without 15 mg/ml Avicel, 1 mM MnSO₄, 1 mM ascorbic acid, 30 to 300 U/liter PsVP (1.5 to 15 μg/ml), and 4 μM PoLPMO9A mixed with 100 mM malonate buffer (pH 4.5). For each degradation reaction, 200 μl was sampled at regular intervals (every 6 h) and mixed with methanol (pH 2.0; final concentration, 80% [vol/vol]) to stop the reaction. The samples were then filtered (0.22-μm pore size) before high-performance liquid chromatography (HPLC) analysis. MALDI-TOF/TOF MS analysis of Avicel incubated with PoLPMO9A and PsVP in the presence of Avicel cellulose and lignin-derived aromatic compound was performed with a mixture containing 15 mg/ml Avicel, 4 μM PoLPMO9A, 500 μM sinapic acid, 1,000 μM Asc, and malonate buffer (100 mM; pH 4.5) at 30°C. Two lignin model dimers (β-O-4 and 5-5′) and two lignin polymers (kraft lignin and natural lignin) were used to investigate the lignin degradation mediated by a LPMO-driven peroxidase reaction. For lignin dimers, the reactions were performed for 48 h with a reaction mixture containing dimer substrates (500 μM), 1 mM MnSO₄, 1 mM ascorbic acid, approximately 30 to 300 U/liter PsVP (approximately 1.5 to 15 μg/ml), 4 μM PoLPMO9A, and 100 mM malonate buffer (pH 4.5). For lignin polymers, the reactions were carried out using 100 mM malonate buffer (pH 4.5) containing 100 mg lignin substrates, 1 mM MnSO₄, 0.5 mM ascorbic acid, 150 U/(g substrate) PsVP, and 0.4 mM/(g substrate) PoLPMO9A. In addition, 25 μg/ml tetracycline was added to prevent microbial contamination. To maintain the reaction, 0.5 mM ascorbic acid was added every 24 h. Positive-control experiments used to evaluate the activity of class II lignin-degrading peroxidase reactions were performed with 150 μM H₂O₂ and 100 U/liter PsVP for degradation of lignin-derived aromatic compounds. For degradation of lignin polymers, 150 μM H₂O₂ was added every 24 h in the positive-control experiments.

The concentrations of lignin-derived aromatic compounds and lignin dimers were determined by HPLC using standard calibration curves for each compound (shown in Table S1 in the supplemental material) (61). To maintain the concentration of each compound prior to HPLC analysis, the samples was mixed with methanol (pH 2.0; final concentration, 80% [vol/vol]) to stop the reaction. Samples (10 μl) were subjected to HPLC analysis with a C18 column (Waters) (4.6 by 250 mm) under conditions of a flow rate of 1 ml/min with methanol–0.1% formic acid (65:35) as the eluent at 40°C. The UV detector operated at λ_max of lignin-derived aromatic compounds and lignin dimers seen in Table S1. The degree (percentage) of degradation of the substrate was calculated using the following formula: degree (percentage) of degradation = [(C_i − C_t)/C_i] × 100% (where C_i represents the initial concentration of the substrate and C_t represents the concentration of the substrate along time). All the results included in Fig. 2 and 3 represent the means and standard deviations of data from three replicates (independent biological samples). Means and standard deviations were calculated using OriginPro 8.0.

Analysis of the products of lignin degradation by PoLPMO9A and PsVP.

The oxidation products of lignin dimers treated with PoLPMO9A-driven PsVP were further analyzed via LC-electrospray ionization (ESI)-MS. The MS spectra were acquired with an Agilent Ion Trap XCT mass spectrometer (Agilent, USA) equipped with electrospray ionization coupled with the HPLC–photodiode-array detection (DAD) system, and initial separation of oxidation products was performed using the same HPLC protocol as described previously. The products of oxidation of lignin dimers by PoLPMO9A-driven PsVP were measured in positive/negative-ion mode, and the electrospray level was set to +3,500 V. Dry gas flow was set to 10 liters/min⁻¹ with a temperature of 300°C, and the nebulizer pressure was set to 40 lb/in² (62). To verify the degradation of kraft lignin and natural lignin, the degradation products were washed with H₂O to remove soluble components and then the residual was freeze-dried for Py-GC/MS analysis. Fast pyrolysis tests were performed using a Pyroprobe 5200 analytical pyrolyzer (CDS Analytical Inc.) at 500°C for 60 s. The pyrolysis volatiles were analyzed using a gas chromatograph (Agilent 7890 A) equipped with a mass spectrometer (5975 CMSD) and HP-5 MS column. The chromatograph program was 1 min isothermal at 40°C, followed by increases at 6°C/min to 300°C, with a final hold at 300°C for 10 min. The MS was performed under 70 eV electron ionization (EI) conditions, with an m/z range of 40 to 400. The pyrolysis products were identified on the basis of the reported literature and the NIST mass spectrum library (63–66). All the results included in Table S3 and Table S4 represent the means and standard deviations of data from three replicates (independent biological samples). Means and standard deviations were calculated using OriginPro 8.0.

Adsorption experiments.

The adsorption of LPMO on the cellulose substrate was investigated as described previously (67). The PoLPMO9A concentrations used in adsorption experiments ranged from 0.0148 to 0.37 mg/ml. Eppendorf tubes containing PoLPMO9A protein, the Avicel substrate (15 mg/ml), and 100 mM malonate buffer (pH 4.5) were incubated in a rotating shaker at 0°C for 180 min. After the incubation, samples were centrifuged at 12,000 rpm for 10 min and passed through a 0.22-μm-pore-size filter (Millipore). The protein concentration in filtrate was quantified using the Bradford method with bovine serum albumin as a standard (53). The level of adsorbed enzyme (in milligrams) was calculated using the following formula: adsorbed LPMO on Avicel (milligrams) = m_i − m_f (where m_i represents the amount of initial PoLPMO9A and m_f represents the amount of free PoLPMO9A). All the results represent means and standard deviations of data from three replicates (independent biological samples). Means and standard deviations were calculated using OriginPro 8.0.