Oxidation of a non-phenolic lignin model compound by two Irpex lacteus manganese peroxidases: evidence for implication of carboxylate and radicals

Background Manganese peroxidase is one of the Class II fungal peroxidases that are able to oxidize the low redox potential phenolic lignin compounds. For high redox potential non-phenolic lignin degradation, mediators such as GSH and unsaturated fatty acids are required in the reaction. However, it is not known whether carboxylic acids are a mediator for non-phenolic lignin degradation. Results The white rot fungus Irpex lacteus is one of the most potent fungi in degradation of lignocellulose and xenobiotics. Two manganese peroxidases (IlMnP1 and IlMnP2) from I. lacteus CD2 were over-expressed in Escherichia coli and successfully refolded from inclusion bodies. Both IlMnP1 and IlMnP2 oxidized the phenolic compounds efficiently. Surprisingly, they could degrade veratryl alcohol, a non-phenolic lignin compound, in a Mn2+-dependent fashion. Malonate or oxalate was found to be also essential in this degradation. The oxidation of non-phenolic lignin was further confirmed by analysis of the reaction products using LC–MS/MS. We proved that Mn2+ and a certain carboxylate are indispensable in oxidation and that the radicals generated under this condition, specifically superoxide radical, are at least partially involved in lignin oxidative degradation. IlMnP1 and IlMnP2 can also efficiently decolorize dyes with different structures. Conclusions We provide evidence that a carboxylic acid may mediate oxidation of non-phenolic lignin through the action of radicals. MnPs, but not LiP, VP, or DyP, are predominant peroxidases secreted by some white rot fungi such as I. lacteus and the selective lignocellulose degrader Ceriporiopsis subvermispora. Our finding will help understand how these fungi can utilize MnPs and an excreted organic acid, which is usually a normal metabolite, to efficiently degrade the non-phenolic lignin. The unique properties of IlMnP1 and IlMnP2 make them good candidates for exploring molecular mechanisms underlying non-phenolic lignin compounds oxidation by MnPs and for applications in lignocellulose degradation and environmental remediation. Electronic supplementary material The online version of this article (doi:10.1186/s13068-017-0787-z) contains supplementary material, which is available to authorized users.


Background
Lignocellulose is a renewable but recalcitrant resource for biofuels and bio-based chemicals [1]. In addition to cellulose and hemicellulose, lignin is one of the major components of lignocellulose. The complex lignin network contains phenolic and non-phenolic lignin substructures, with the latter constituting the major part. The white rot fungi (WRF) are regarded to be the best lignocellulose degraders, whose enzymatic systems have hence been an object of extensive studies [2]. WRF produce a range of lignin-modifying enzymes, which include lignin peroxidase (LiP, EC 1. 11 [3]. In addition, free radicals of different types such as hydroxyl radical (OH·), carboxylate anion radical (CO ·− ), and superoxide radical (O 2 ·− ) generated by WRF are also implicated in lignocellulose depolymerization [4]. Among the lignin-modifying enzymes, LiP, VP, and DyP are capable of directly oxidizing non-phenolic lignin model compounds such as veratryl alcohol (VA), whereas MnP and Lac do not have this property [5]. Interestingly, however, MnP and Lac appear to be the most abundant lignin-modifying enzymes for many WRF [3]. This suggests that MnP and Lac may use an alternative mechanism(s) to oxidize the high redox potential non-phenolic lignin moiety.
It is well known that MnP can oxidize Mn 2+ to Mn 3+ , which forms chelates with organic acids to directly attack the low redox potential phenolic lignin. During this process, unstable free radicals are formed, which tend to disintegrate spontaneously [6]. The chelated Mn 3+ ions can also react with a certain co-oxidant (or mediator) to generate reactive radicals that can depolymerize the high redox potential non-phenolic lignin. Unsaturated fatty acids (UFA) and their lipid derivatives are such mediators that can be peroxidized to form highly reactive acyl and fatty acid peroxyl radicals acting on the non-phenolic lignin [7][8][9][10][11]. Other mediators such as glutathione (GSH) may also be involved in the formation of thiyl radicals, which are thought to be also involved in the degradation of recalcitrant compounds [12,13]. However, whether organic acids, particularly those excreted by fungi, are implicated in degradation of high redox potential nonphenolic lignin and xenobiotics has never been clearly demonstrated.
The white rot fungus Irpex lacteus has a strong potential in biopretreatment of lignocellulose as well as in biodegradation of xenobiotic compounds. I. lacteus appears to produce MnP as the main ligninolytic enzyme under tested conditions [14,15]. I. lacteus CD2 is a strain isolated from Shennong Nature Reserve (Hubei, China) with outstanding capability in degrading lignin and dyes. Although a few MnPs have been purified from the I. lacteus cultures, it is not known how these enzymes are involved in destructing lignin and xenobiotics [16,17]. Herein, we expressed two MnP genes from I. lacteus CD2 in Escherichia coli and successfully refolded them from inclusion bodies. We showed evidences that MnP-oxidized Mn 3+ may chelate with a carboxylic acid and form radicals, which are further implicated in degradation of non-phenolic lignin and high redox potential dyes.

Results and discussion
Gene cloning and sequence analysis of IlMnP1 and IlMnP2 The MnPs of I. lacteus CD2 have been reported to play an important role in the biological pretreatment of lignocellulose and decolorization of synthetic dyes and even simulated textile wastewater [15]. However, the corresponding mechanism involved in lignin depolymerization and dyes decolorization was unclear. In the present study, two MnP genes (GenBank accession number KX620478 and KX620479), 1684 and 1622 bp, were identified in the genome of I. lacteus CD2 (Additional file 1), and their respective cDNAs were successfully obtained from the culture grown on BM medium. The IlMnP1 and IlMnP2 were interrupted by 11 introns and 10 introns, giving two open reading frames (ORFs) of 1077 and 1080 bp, respectively (Additional file 1). Deduced IlMnP1 and IlMnP2 contained 358 and 359 amino acid residues and harbored a signal peptide of 18 and 21 residues, respectively. Similar regulatory elements including TATA box, CAAT motif, CreA-and NIT2-binding sites, putative heat-shock element (HSE), and xenobiotic-responsive element (XRE) were discovered in the upstream region of both genes (Additional file 1). Carbon catabolite repression mediated by CreA or its orthologs was widely found both in ascomycetes [18] and basidiomycetes [19]. The presence of CreA-binding sites implied that the expression of the two MnP genes might be repressed by glucose.

Refolding and purification of IlMnP1 and IlMnP2 expressed in E. coli
Pichia pastoris and Trichoderma reesei, the two popular microbial systems for large-scale production of commercial enzymes, were firstly used as the expressing host but the attempts to express IlMnP1 and IlMnP2 in these two microbes failed. E. coli was then chosen to express these two enzymes. Both IlMnP enzymes accumulated exclusively in the inclusion bodies, as had been reported previously for other MnP, VP, and LiP [20,21]. The inclusion bodies were then solubilized using urea as described previously [20]. Multiple factors including pH, hemin, urea, GSSG, and refolding time are all critical for the successful refolding of peroxidases. By using a fast screening method with 96-well microplates, the optimum pHs for refolding of both enzymes were determined to be pH 9.5 (Fig. 1a), which were the same as that for a VP from Pleurotus eryngii [22]. Alkaline pHs were favorable for the formation of thiolate anion, which was essential for the formation of disulfide bridges [22]. Note that both MnPs were predicted to have four disulfide bridges. Different urea concentrations were required for the maximal yield of active IlMnP1 (0.2 M) and IlMnP2 (0.5 M) in refolding (Fig. 1b). The requirements of MnPs from I. lacteus CD2 for urea were much lower than other Class II fungal peroxidases (up to 2 M) [21]. The reducing agents GSSG and DTT were also essential for the formation of disulfide bridges. As shown in Fig. 1c, the optimal GSSG/ DTT ratios for the MnPs were 5:1 (0.5 mM GSSG versus 0.1 mM DTT). Although hemin was not necessary for the refolding of other Class II fungal peroxidase or the horseradish peroxidase, it was required for the refolding of IlMnP1 and IlMnP2 at an optimal concentration of 10 μM (Fig. 1d). Over the time course, the refolding of IlMnP2 significantly increased from 10 to 20 h to reach a plateau, while that of IlMnP1 decreased instead from 10 h (Fig. 1e).
Large-scale refolding of the IlMnP1 and IlMnP2 was conducted under the optimized conditions (0.5 mM GSSG, 0.1 mM DTT, 10 μM hemin, 5 mM CaCl 2 , and 0.1 mg/mL protein, 0.5 M urea for IlMnP1 or 0.2 M urea for IlMnP2, pH 9.5) for 10 h at 15 °C. The refolded proteins were further purified by anion exchange. Finally, yields of 28.2 mg and 13.3 mg of functional IlMnP1 and IlMnP2, respectively, per liter culture were obtained. Both enzymes showed a single band on SDS-PAGE gels, corresponding to the calculated molecular masses (Additional file 2). Moreover, the enzymes had an absorbance peak at 409 nm ( Fig. 2), indicating that each MnP harbors a heme group [23]. The Rz (A 407 /A 280 ) ratios of IlMnP1 and IlMnP2 were 1.0 and 2.4, respectively.

Optimal pH and temperature of IlMnP1 and IlMnP2
The optimal pHs of recombinant IlMnP1 and IlMnP2 were both pH 4.0 (Additional file 3a). When the pH was above 6.5, no activity was detected for both enzymes. This character was similar to that of native MnPs from I. lacteus strains CD2 (pH 3.0-6.0) and Fr. 238 (pH 3.0-7.6) and other fungi, which are all acidic MnPs ( Table 1). The two MnPs varied in pH stability (Additional file 3b). At neutral pH, the IlMnP2 retained much more residual activity than IlMnP1. Interestingly, most native or recombinant MnPs from I. lacteus ever reported exhibit remarkable stability at neutral pH, while one MnP from Phanerochaete chrysosporium was inactive at near neutral pH (6.5) [24]. The optimal temperatures of IlMnP1 and IlMnP2 were both 60 °C (Additional file 3c). However, at 60 °C both enzymes quickly lost their activity within 10 min (Additional file 3d). The thermostability of the two recombinant IlMnPs were not as good as a natively purified MnP from I. lacteus CD2: the native I. lacteus CD2-MnP retained 93.2% of the initial activity after 1 h of incubation at 40 °C. At this temperature, IlMnP2 retained 80.5% activity, while IlMnP1 had only 13.1% left after 1 h of incubation [15]. This weakness in thermostability might be ascribed to the lack of glycosylation during heterologous expression in E. coli [25].

Biochemical analysis of IlMnP1 and IlMnP2 on Mn 2+ and phenolic lignin model compounds
The K m values of IlMnP1 and IlMnP2 for Mn 2+ were 193.8 and 152.2 μM, respectively, higher than those of native MnPs (17-49 μM) ( Table 1). The k cat values of IlMnP1 and IlMnP2 were 7.1 and 6.6 s −1 , respectively. The structures and maximal absorbance wavelengths of the substrates (phenolic, non-phenolic lignin model compounds, and dyes) used in this study are listed in Table 2. IlMnP1 and IlMnP2 could oxidize two  phenolic substrates DMP and guaiacol as well as the substrate ABTS, with the specific activities significantly higher in the presence of Mn 2+ (   [16]. The oxidation of phenolic substrates by MnPs was thought to be through one-electron oxidation involving the chelated Mn 3+ ions [6].

Degradation of a non-phenolic lignin model compound by IlMnP1 and IlMnP2
Surprisingly, the two MnPs could also oxidize the nonphenolic substrate VA albeit only in the presence of Mn 2+ (Fig. 3a). No MnP has been reported previously to have VA-oxidizing ability, which was thought to be a unique catalytic feature of high redox potential peroxidases such as LiP and VP. These VA-oxidizing enzymes commonly have a tryptophan residue involved in VA binding near the heme-binding site [26]. Interestingly, neither IlMnP1 nor IlMnP2 bears such a characteristic tryptophan (aspartate for IlIMnP1 and alanine for IlIMnP2 at the corresponding position instead, Additional file 4), excluding the possibility that IlMnP1 and IlMnP2 are LiPs or VPs. The unusual catalysis of VA suggested that another mechanism must be involved in oxidation of VA by IlMnP1 and IlMnP2.
In order to confirm that IlMnP1 and IlMnP2 could oxidize the non-phenolic lignin model compound VA, the reaction products were further analyzed by HPLC as well as LC-MS/MS. A peak corresponding to veratraldehyde was clearly detected in reactions in the presence of IlMnP1 and IlMnP2, Mn 2+ , and malonate (Fig. 3a) or oxalate (Fig. 3b) but not with acetate, citrate, lactate, or succinate (Additional file 5).  (Fig. 4a). These ions could also be observed in the MS/MS analysis of IlMnP2-VA reaction products, confirming that IlMnP2 could indeed convert VA into veratraldehyde (Fig. 4) [27].
The catalysis depends on the presence of both Mn 2+ and a specific carboxylic acid since VA was not degraded when Mn 2+ was absent (Fig. 3a) or if malonate was replaced by acetate, citrate, lactate, or succinate (Additional file 5). Acetate and succinate could not form complexes with Mn 3+ [28]; therefore, no oxidation product was observed in the acetate or succinate buffer. Mn 3+ can form chelates with the rest four organic acids. Mn 3+ -lactate/tartrate complexes were reported  [28]. Therefore, we hypothesized that Mn 3+ -lactate/citrate complexes was more apt to react with H 2 O 2 than malonate/ oxalate. However, this phenomenon was not detected for the malonate or oxalate buffer [28]. Moreover, it was reported that the carbon-centered radical and superoxide radical were generated from the oxidation of malonate/ oxalate by Mn 3+ [29,30]. The VA-oxidizing activities of IlMnP1 and IlMnP2 in malonate were higher than those in oxalate (Fig. 3b), and oxalate was also reported to be ineffective in supporting VA oxidation by MnP from Lentinus edodes [31]. These clearly indicated that both Mn 2+ and the carboxylate play an indispensable role in degrading the non-phenolic lignin model compound by IlMnP1 and IlMnP2. GSH and UFA have been reported to mediate oxidation of non-phenolic lignin compounds by MnPs through generation of highly active thiyl and fatty acid peroxyl radicals, respectively [6,13]. Note that carboxylic acids can also be oxidized by chelated Mn 3+ to generate radicals [6,29]. We also noticed that the extent of VA oxidation by IlMnP1 and IlMnP2 improved when enzyme loading increased from 0.05 to 0.25 and then to 0.5 U/mL, particularly for IlMnP2 (Fig. 5a). Besides, the oxidation product veratraldehyde steadily increased with a linear relationship to the concentrations of VA (Additional file 6). However, whether the radicals generated in the MnP-Mn 2+ -carboxylic acids system are able to attack the high redox potential non-phenolic lignin is not known from previous literature studies. Since Mn 2+ and a certain carboxylate (malonate and oxalate) are the two necessary components needed for IlMnP1 and IlMnP2 to degrade non-phenolic lignin compounds, it is now reasonable to infer that the ability of IlMnP1 and IlMnP2 to oxidize VA was actually through the action of radicals, which were generated by the reactions of MnPoxidized Mn 3+ with malonate or oxalate. To gain some insights of VA oxidation by the two IlMnPs, the reactions in the malonate buffer were performed in the presence or absence of SOD (at a final concentration of 3000 U/ mL), which is a commonly used scavenger for superoxide radical [32]. SOD had no inhibitory effect on the formation of Mn 3+ (for IlMnP1, 1048 and 1053 U/L in absence and presence of SOD, respectively; for IlMnP2, 965 and 1000 U/L in absence and presence of SOD, respectively) but partially inhibited the oxidation of VA by IlMnP1 (by 25.3%) and IlMnP2 (by 23.9%) (Fig. 5b). This indicated that the superoxide radical is at least partially responsible for VA oxidation by the two MnPs in presence of a certain carboxylic acid. Interestingly, both malonate and oxalate are known to be acids excreted by saprophytic fungi including I. lacteus and P. chrysosporium [8,28]. Our results suggest that I. lacteus may use its MnPs with a particular organic acid(s) it excretes to co-operate in degrading the more recalcitrant lignin.
Interestingly, while the pH optimum for LiP, VP, and DyP in oxidizing VA is pH 3 or lower [5,26,33], IlMnP1 and IlMnP2 exhibited VA-oxidizing ability at pH 5, which is also optimal for the mostly used T. reesei cellulases [34] and similar to those of many other acidic plant cell wall polysaccharides degrading enzymes [35,36]. MnP, in the presence of Mn 2+ and a carboxylic acid mediator such as malonate, may hence be used to formulate enzyme cocktails with cellulase and hemicellulase to simultaneously deconstruct lignin, cellulose, and hemicellulose. Besides the application potential in lignocellulose degradation, IlMnP1 and IlMnP2 may also serve as good candidates for further investigating the molecular mechanisms underlying non-phenolic lignin depolymerization by MnPs.

Application potential of IlMnP1 and IlMnP2 in decolorizing dyes with different structures
IlMnP1 and IlMnP2 are capable of directly or indirectly degrading phenolic and non-phenolic lignin compounds with varying structures, which enlightens us to explore if they also have the ability to decolorize dyes for environmental remediation. A purified MnP from I. lacteus CD2 has been reported to be able to efficiently decolorize different types of dyes [15]. IlMnP1 and IlMnP2 also exhibited strong ability to decolorize a broad range of dyes including the azo dyes (RBV5R and RB5), anthraquinone dye (RBBR), indigo dye (IC), and triphenylmethane (MG) in the presence of Mn 2+ and malonate (Fig. 5). The decolorization of RBV5R and IC was the fastest: above 85% of the dyes (50 mg/L) could be decolorized by the enzymes within 1 h (Fig. 6a, d). In contrast, the degradation of RBBR and MG was much slower; more than 90% of the dyes could be decolorized after 5 h of incubation (Fig. 6c, e). Nonetheless, the decolorization of MG by IlMnP1 and IlMnP2 was much more effective than that by the purified MnP from I. lacteus CD2 (32% decolorized after 36 h of incubation [15]). The degradation of RB5 was the slowest: 31.9 and 25.4% were decolorized by IlMnP1 and IlMnP2, respectively, within 10 h (Fig. 6b). RB5 is considered a specific substrate for VP but not oxidized by the MnP of P. chrysosporium in a tartrate buffer [37]. Note that the rate of dye decolorization was significantly reduced in the absence of Mn 2+ for all dyes. These together support the notion that, like lignin degradation, Mn 2+ and a specific carboxylate such as malonate are important constituents for efficient dye decolorization by the two MnPs from I. lacteus CD2.

Conclusions
In this study, two manganese peroxidase genes were cloned from the white rot fungus I. lacteus CD2. By optimizing a variety of parameters, the E. coli-expressed IlMnP1 and IlMnP2 were successfully refolded from inclusion bodies. The recombinant IlMnP1 and IlMnP2 could oxidize a series of phenolic and even non-phenolic lignin model compounds substrate VA. Mn 2+ and a certain carboxylate (malonate or oxalate) are the two indispensable components in enzymatic degradation of the non-phenolic lignin. It is proposed that radicals such as superoxide radical formed in this carboxylate buffer system are at least partially involved in degrading these high redox potential lignin compounds. Besides, IlMnP1 and IlMnP2 could also decolorize dyes of four different types, whose efficiency also depended on the presence of Mn 2+ . In summary, we demonstrated that the degradation of non-phenolic lignin by MnP is not restricted to GSH or UFA mediators but can expand to carboxylic acids, which are excreted by fungi as a normal metabolite. The properties of IlMnP1 and IlMnP2 make them ideal candidates for exploring molecular mechanisms underlying lignin deconstruction by MnPs and potential players in formulating efficient enzyme cocktails for lignocellulose degradation and dye decolorization.  Table 2.

Cloning and expression of IlMnP1 and IlMnP2
Irpex lacteus CD2 was grown for 5 days in the basal liquid medium [15]. Total RNA was extracted using the TRIZOL reagent (Invitrogen, Waltham, MA) according to the manufacturer's instructions. The first strand cDNA was synthesized from the total RNA using the TransScript One-Step gDNA Removal and cDNA Synthesis Supermix with oligo (dT) (TransGen). Based on the 5′-and 3′-end sequences of the IlMnP1 and IlMnP2 structural genes, the MnP genes devoid of the sequences encoding the signal peptides were amplified with genespecific primers (as shown in Additional file 7). The PCR products were T-A ligated into pEASY-T3 (TransGen) and then transformed into the E. coli Trans1-T1 to obtain pEASY-T3-IlMnP1 and pEASY-T3-IlMnP2.
The recombinant plasmids pEASY-T3-IlMnP1 and pEASY-T3-IlMnP2 were double-digested with BamHI/NotI and BamHI/XhoI, respectively, gel purified, and ligated into the pre-digested pET-28a(+) to obtain pET28a-IlMnP1 and pET28a-IlMnP2, and individually transformed into E. coli BL21 (DE3) competent cells. The cells harboring pET28a-IlMnP1 or pET28a-IlMnP2 were pre-cultured in LB medium supplemented with 50 μg/mL of kanamycin at 37 °C overnight with shaking at 200 rpm and used as the inocula of 200 mL LB medium. The cultures were grown at 37 °C for 3 h, followed by the addition of isopropyl-β-d-thiogalactoside (IPTG) to a final concentration of 1 mM for 4-h induction.
After induction, the cells were harvested by centrifugation. The pellets were re-suspended in 50 mM Tris-HCl, 10 mM EDTA, and 5 mM DTT (pH 8.0). Lysozyme (Amresco, Solon, OH) was then added to a final concentration of 2 mg/mL and the cells were incubated on ice for 1 h. Then, 20 μL of DNase I (TransGen) was added and the incubation was continued on ice for 30 min. Subsequently, the cells were centrifuged at 12,000g for 30 min at 4 °C. No MnP activity could be detected from the supernatants. The cell debris was washed with 20 mM Tris-HCl, 1 mM EDTA, and 5 mM DTT (pH 8.0) twice, followed by incubation in 50 mM Tris-HCl, 8 M urea, 1 mM EDTA, and 1 mM DTT (pH 8.0) on ice for 1 h.
To optimize the parameters for recovering active enzyme from inclusion bodies, the refolding was performed in various conditions in a 200 μL volume using 96-well plates at 15 °C. A range of parameters including concentrations of urea, GSSG, and hemin and pH were investigated, while the concentrations of enzyme, EDTA, and DTT were kept constant during the refolding. The efficiency of refolding was indicated by the MnP activity. Based on the fast plate-screening result, large-scale refolding of MnPs was performed using the respective optimum parameters. After refolding, the crude enzymes were centrifuged at 12,000g for 10 min at 4 °C and the insoluble fractions were discarded. The supernatants containing the refolded MnP were concentrated through a 10 kDa cut-off centrifuge filter, followed by dialysis against 20 mM phosphate buffer, pH 6.0. The crude enzymes were further purified by a HiTrap Q HP anion exchange column (GE Health, Fairfield, CT) pre-equilibrated with the same phosphate buffer. The proteins were eluted with a linear gradient of 0-1.0 M NaCl, and fractions containing active enzymes were pooled.

Biochemical characterization of IlMnP1 and IlMnP2
The refolded IlMnP1 and IlMnP2 were first subjected to UV-visible spectroscopic analysis in the range of 230-800 nm in the 20 mM malonate buffer (pH 5.0). The MnP activity was measured by monitoring the oxidation of ABTS (ε 420 = 36,000 M −1 cm −1 ) at 420 nm, in a buffer containing 50 mM malonate, 1 mM ABTS, 1 mM MnSO 4 , and 0.1 mM H 2 O 2 (pH 5.0 and 25 °C). For the Mn 2+ -independent activity assay, MnSO 4 was omitted. One unit (U) of enzyme activity was defined as the amount of enzyme that oxidizes 1 μmol of ABTS per min at 25 °C [30]. For kinetic studies, the reactions were performed in the 50 mM malonate buffer (pH 5.0) at 25 °C using 10-4000 μM Mn 2+ (in the presence of 0.1 mM H 2 O 2 ) as the substrate by monitoring the formation of Mn 3+ -malonate complexes (ε 270 = 11,590 M −1 cm −1 ) at 270 nm [15]. The non-linear least square fitting method was used to calculate the K m , k cat , and k cat /K m parameters of the recombinant IlMnP1 and IlMnP2 using the Graph-Pad Prism 5 software.
To determine the pH optimum, the MnP activity on ABTS was determined in the 20 mM malonate buffer at a pH ranging from 3.0 to 7.0 at 25 °C. For temperature optimum, the enzymatic activity was measured in the 20 mM malonate buffer (pH 5.0) at a temperature from 20 to 80 °C. To evaluate the pH stability, IlMnP1 and IlMnP2 were individually incubated at different pH levels (3.0-7.0) for 1 h, and the residual activities were assayed as described above. For thermostability, IlMnP1 and IlMnP2 were incubated at 40-60 °C for 1 h with samples taken for activity measurement periodically. The residual activities were measured at its optimum pH and temperature.
The substrates specificities of IlMnP1 and IlMnP2 were studied for the oxidation of four different substrates ABTS, DMP, guaiacol, and VA in 50 mM pH 5.0 malonate and 0.1 mM H 2 O 2 with or without 1 mM MnSO 4 . Activities were calculated using absorption coefficients at the corresponding wavelengths.

Oxidation of non-phenolic lignin model compounds by IlMnP1 and IlMnP2
VA was used in evaluating the abilities of IlMnP1 and IlMnP2 for degradation of the non-phenolic lignin compound. The degradation of VA was performed in 50 mM malonate buffer (pH 5.0) containing 1 mM VA, 1 mM MnSO 4 , 0.1 mM H 2 O 2 , and 0.5 U/mL IlMnP1 or IlMnP2. In some reactions, the malonate buffer was changed to another carboxylate (acetate, oxalate, citrate, lactate, or succinate) buffer (pH 5.0) or MnSO 4 was omitted from the reaction. The effect of VA concentration (0.05-1 mM) on oxidation was analyzed in the malonate buffer (pH 5.0) for 48 h with 0.5 U/mL IlMnP1 or IlMnP2 in the presence of 1 mM Mn 2+ . The effect of enzymes loading and superoxide dismutase (3000 U/mL) on VA oxidation by IlMnP1 or IlMnP2 was analyzed in the malonate buffer (pH 5.0) for 48 h in the presence of 1 mM Mn 2+ . The ability of Mn 3+ formation was evaluated by the oxidation of ABTS. The reaction proceeded at 30 °C for 48 h, and then the reaction products were analyzed by HPLC using a reversed phase C 18 -column (Eclips XDB-C 18 , 4.6 mm × 250 mm, 5 μm). The elution condition was 0% Acetonitrile (ACN), 4 min; 0-60% ACN, 10 min; 60-100% ACN, 1 min; and 100% ACN, 5 min at a flow rate of 0.8 mL/min. The elution peaks were monitored at 310 nm. In order to confirm veratraldehyde as the oxidation product, LC-MS was also performed by coupling a Nexera UHPLC system to an AB-SCIEX 5600 Triple TOF mass spectrometer in positive and high-sensitivity mode. For LC analysis, the column, mobile phase, detection, and flow rate were identical to those for HPLC analysis described above. The elution program was as follows: 0-60% ACN, 10 min; 60-100% ACN, 1 min; and 100% ACN, 5 min. For MS analysis, the parameters were set as ion source gases GS1, GS2, and curtain gas were 55, 55, and 35 psi, respectively, temperature was 600 °C, and ion spray voltage floating was at 5500 V.

Decolorization of dyes by IlMnP1 and IlMnP2
Five dyes of different structures were used to evaluate the decolorization capability of IlMnP1 and IlMnP2. The reactions were carried out at 30 °C in a total volume of 200 μL containing 50 mM malonate buffer (pH 5.0), 0.1 mM H 2 O 2 , 0.25 U/mL IlMnP1 or IlMnP2, and 50 mg/L of dye, with or without 1 mM Mn 2+ . During the incubation, the color changes were periodically detected by measuring the optical density (OD) at 556 nm for RBV5R, 596 nm for RB5, 600 nm for RBBR, 610 nm for IC, and 640 nm for MG. The rate of decolorization was then calculated using the following formula: decolorization (%) = [(A i − A t )/A i ]×100, where A i and A t are the absorbance at the initial and given stages. Authors' contributions XZ, XYS, and BY conceived and designed the experiments. XQ and XHS performed the experiments. XQ, HH, and YB analyzed the data. XQ and XYS wrote the manuscript. YW and HL reviewed and revised the manuscript. All authors read and approved the final manuscript.