For industrial production, the temperature for vegetable growth of L. decastes is ~23°C. After ~60 days of cultivation, the surface mycelia were scratched and the temperature was adjusted to 16–18°C, 90%–95% humidity. The fruiting bodies were harvested 25–30 days after scratching. The dikaryotic strain LRG-d1 was isolated from the fruiting body of industrially cultivated L. decastes (Figure 1). The Basidiomycetes spores were collected from the fruiting body of strain LRG-d1. The spores were spread on the PDA plates by serial dilution. The monokaryotic strain without clamp connection was isolated and verified by micro-examination. The mating compatibility between different monokaryotic strains was determined by confrontation culture. Monokaryotic strains that could form clamp connections were selected for further analysis as previously described (Li et al., 2022). Three monokaryotic strains were cultivated on a PDA plate covered with cellophane as previously described (Li et al., 2022). After spreading all the plate, the mycelia were scraped from the cellophane, frozen in liquid nitrogen, and stored at −80°C for genome sequencing and RNA-Seq.

Genome DNA was extracted from mycelia of monokaryotic strain LRG-1 using the NucleoBond HMW DNA kit (Macherey-Nagel, Düren, Germany) according to the manufacturer’s instructions. The Nanopore library was constructed using Oxford Nanopore LSK-109 kit, and the Nanopore library was sequenced on the PromethION platform by Benagen Technology Co., Ltd. (China, Wuhan). After quality filtered using Filtlong v0.2.0 (Wick, 2017), a total of 6.55 Gb and 9.19 Gb clean raw data were generated for strain LRG-d1-1 and LRG-d1-5, respectively. Illumina paired-end sequencing was sequenced on an Illumina NovaSeq 6000 platform under 150 bp mode in the same company. The raw fastq data were filtered using fastp software, and 4.86 Gb, 3.51 Gb, and 5.27 Gb raw data were generated after filtration for strain LRG-d1-1, LRG-d1-5, and LRG-d1-2, respectively.

Raw Oxford Nanopore reads were assembled using flye v2.8.3-b1695 and NextDenovo v2.4.0 software with default parameters. The data reads were also assembled using necat with parameters: GENOME_SIZE = 48 m and CNS_OUTPUT_COVERAGE = 70. The assembled contig was polished through two iterations using Racon¹ with Nanopore reads and default parameters. The polished contigs were further polished with Pilon v1.24 through two iterations using filtered Illumina reads (Walker et al., 2014). GC content = (G + C)/(A + T + G + C).

Genes were predicted with Augustus software using laccaria_bicolor models (Stanke et al., 2006). Genes were predicted using GeneMark-ES v 4.69 for self-training prediction using default parameters (Ter-Hovhannisyan et al., 2008). RNA-Seq data were mapped using stringtie software (Pertea et al., 2015) and the results were used for model training with Augustus software, the genes were also predicted using the trained model based on RNA-Seq transcripts. The three prediction genes were integrated using EVidenceModeler software (Haas et al., 2008). Functional annotations of predicted protein-coding sequences (CDSs) against EggNGOmapper, Pfam, and SwissProt were performed as previous description (Yu et al., 2022). CAZymes of the L. decastes genome were annotated using dbcan version v3.0.2 software with Hmmer search engine and default parameters as previously described (Yu et al., 2022). The signal peptide of a protein was predicted using SignalP v5.0b software (Almagro et al., 2019). The trans-membrane structure was predicted using TMHMM v2.0c (Chen et al., 2003). Protein that has signal peptide without trans-membrane structure was secreted protein.

The pairwise average nucleotide identity (ANI) analysis was performed using fastANI software (Jain et al., 2018). Collinearity analysis was performed using MCScanX software (Wang et al., 2012). Orthogroups analysis was using OrthoFinder software (Emms and Kelly, 2019). Divergent time analysis and gene family evolution were performed using Bayesian software packages MCMCTree and CAFE v5.0 software, respectively, (Yang, 2014; Mendes et al., 2020).

The mating-type loci of L. decastes were analyzed using sequence alignment with homeodomain (HD) genes and pheromone receptor genes (STE) genes from Hypsizygus marmoreus and Pleurotus giganteus as the query sequences (Wang et al., 2021b; Yu et al., 2022). The functions of the identified genes were further confirmed using Blastp tools in the NCBI database. Gene cluster structure was visualized using integrative genomics viewer software (Thorvaldsdóttir et al., 2013).

Dikaryotic strain LRG-d1 was inoculated and cultivated on a PDA plate. Three mycelial plugs (8 mm diameter) were cut from the margin of seed agar cultures and inoculated into 50 mL PDB media (extract from 200 g potato, 20 g dextrose) and 50 ml lignin-rich media (50 mL PDB media add 1 g sawdust, 1 g cottonseed hull, and 1 g bran, respectively) in 250 ml triangular flasks, respectively. Submerged cultivation of L. decastes mycelia was performed at 25°C in darkness at 150 rpm. Three replicates were performed for each condition.

After 10 days of cultivation, the culture was filtered using 6-layer absorbent cotton gauze. The filters were centrifuged at 10,000 rpm for 20 min. The secreted proteins in the supernatant were extracted using phenol extraction coupled with ammonium acetate precipitation as described by Isaacson et al. (2006). Protein pellets were re-dissolved in 4% SDS buffer (in 100 mM Tris–HCl buffer, pH 7.6) and digested using trypsin with STrap methods as described by Zougman et al. (2014). LC-MS analysis was performed in DDA mode as in our previous description without changing (Lin et al., 2022), and the chromatographic time is 90 min. Three biological replicates were performed for each group.

Proteins were identified with at least two unique peptides with a false discovery rate lower than 0.05 using MaxQuant software with default parameters (Tyanova et al., 2016a). For protein expression quantification, MaxQuant software was used to calculate the intensity-based absolute quantification (iBAQ) abundances for each protein. Abundance data obtained from MaxQuant software was processed using Perseus software (Tyanova et al., 2016b). Proteins were normalized using sum normalization methods. Visualization was performed using the R program with ggplot2 and heatmap2 packages.

Monokaryotic strain LRG-d1-1, LRG-d1-2, and LRG-d1-5 were isolated from the basidiospore of dikaryotic strain LRG-d1 (Figure 1). Monokaryotic L. decastes strain LRG-d1-2 and LRG-d1-5 were mating compatible strains to strain LRG-d1-1. To comprehensive analyze the genome of L. decastes, the genomes of three monokaryotic strains were sequences.

The genome of the L. decastes monokaryotic strain LRG-d1-1 strain was sequenced using Oxford Nanopore and Illumina sequencing platforms. De novo assembly was performed with Necat software using the long reads generated by the PromethION platform of Oxford Nanopore Technologies. The final genome assembly of strain LRG-d1-1 consists of 15 contigs, which spans 47.72 Mb in total, with the contig N50 size of 3.92 Mb and N90 size of 2.08 Mb (Table 1; Figure 2). The average GC content of the CDSs of L. decastes LRG-d1-1 (50.25%) is similar to that of L. decastes JCM 30590 (50.00%), Lyophyllum semitale D66 (50.06%) and Lyophyllum shimeji JCM 30591 (50.38%), while lower than that of L. atratum CBS 144462 (51.34%). Repeat annotation revealed that 25.67% (12.25 Mb) of the assembled L. decastes genome comprised repeat elements (Supplementary Table 1). We compared the proportion of repeat elements in seven edible mushrooms, Agaricus bisporus var. burnettii H119_p4 (15.31% in 30.7 Mb genome), Flammulina velutipes 6–3 (12.79% in 38.1 Mb genome), Lentinula edodes L808 (27.23% in 45.9 Mb genome), Pleurotus ostreatus PC15 (9.08% in 34.3 Mb genome), Pleurotus giganteus zhudugu2 (11.36% in 40.0 Mb genome), Stropharia rugosoannulata A15 (17.80% in 47.9 Mb genome), and L. decastes LRG-d1-1. It seems that larger genome has larger repeat elements proportion. Similar to most edible mushrooms, LTR elements were the dominant class of repeat elements. Gypsy/DIRS1 LTR elements accounted for 10.67% of the genome. The genome of L. decastes strain LRG-d1-2 was sequenced using the Illumina platform and the genome of strain LRG-d1-5 was sequenced using Illumina and Nanopore platforms. The final genome assembly of strain LRG-d1-5 consists of 18 contigs, which spans 42.98 Mb in total, with the contig N50 size of 3.53 Mb. The genome size of strain LRG-d1-2 is 40.04 Mb in 2,400 contigs.

Based on a comprehensive strategy of ab initio gene prediction and homology evidence from RNA-Seq transcripts, 14,499 non-redundant gene models were predicted from the genome of L. decastes with an average sequence length of 1,652.2 bp and an average exon number of 6.25 (Table 2; Figure 1). The total length of CDSs was 23.95 Mbp accounting for 50.2% of the total genome. The predicted gene sequence was annotated functionally by sequence alignment against sequences in EggNOGmapper, SwissProt, Pfam, and CAZymes databases. Among the annotated genes, 11,669 and 6,415 genes were classified by the EggNOGmapper database and SwissProt database (Supplementary Table 3), respectively. A total of 8,655 (59.7%) genes were annotated by the Pfam database based on the similarity of protein domains. The completeness of gene prediction was evaluated with 758 BUSCO genes from fungi_odb10, of which 725 BUSCOs (95.7%) were complete (Supplementary Figure 1). Which indicated a high quality of the genome assembly and high fidelity of gene prediction.

The average nucleotide identity (ANI) was analyzed to estimate genomic differences and relatedness between sequenced Lyophyllum strains. As shown in Supplementary Figure 2, four L. decastes strains showed high genomic similarities width ANI value higher than 92%. The results confirmed that these strains are from the same species. The ANI values between different species were lower than 83. The Lyophyllum strains showed no ANI value with strains from the Trichoderma genus (data not shown). The results confirmed the phylogenetic relationship of these strains based on morphological features and ribosomal sequences.

All-against-all comparisons were performed using OrthoFinder to identify orthologs of L. decastes with orthologues in 21 fungi species. Concatenated amino acids from 346 single-copy orthologs were used for phylogenetic reconstruction and species divergence time estimation. Four species from the Lyophyllaceae family (H. marmores, L. atratum, L. decastes, and Termitomyces sp.) were clustered in one clade (Figure 3A), and the results confirmed that taxonomy based on morphology and ITS sequences. We estimated that the divergence of species from the Lyophyllaceae family lineage and Agaricaceae family lineage (A. bisporus) was estimated to occur ~52.34 MYA (Figure 3A). Lyophyllum decastes split from Termitomyces sp. around 40.51MYA. Lyophyllum decastes and L. atratum shared a common ancestor about ~37.32 MYA ago.

For inter-species comparative genomic studies, the synteny of the L. decastes genome and two chromosome-scale assembled genome in Agaricales (A. bisporus and P. ostreatus) were analyzed (Figure 3B). Significant collinearity was detected between the three species. Ten contigs in L. decastes (chr01, chr02, chr03, chr04, chr06, chr07, chr08, chr10, chr11, and chr13) show collinearity with only one chromosome in P. ostreatus and intra-chromosome rearrangements were observed between these chromosomes. Some inter-chromosome rearrangements can also be observed. Three blocks from chr05, chr12, and chr14 show collinearity with chromosome KL198013 in P. ostreatus. Similar collinearity was observed between L. decastes and A. bisporus, only three contigs (chr01, chr05, and chr06) show collinearity blocks with two chromosomes in A. bisporus. The results indicated that genome structure variations in L. decastes mainly occurred inside a chromosome instead of rearrangement between different chromosomes.

Compared with the ancestor of L. decastes and H. marmoreus, gene family expansion and contraction analysis classified 426 and 834 gene families as expanded and contracted, respectively (Figure 3C). Among these, 58 were identified as rapidly/significantly evolving families (p-value < 0.1), including 39 expanded and 19 contracted (Supplementary Table 4). Functional annotation of the 1,194 expanded gene families revealed that they were mainly involved in transposon (10 gene families) and regulation (including kinases, proteins with SNF2-related domain, and Ankyrin repeats). Therefore, the results indicated that regulation might play important role in the physiological differences between H. marmoreus and L. decastes. Intragenome comparisons within the L. decastes genome revealed that no clear whole-genome duplication (WGD) events during evolution (data not shown), which indicated tandem duplication instead of WGD events might play an important role in the expansion of gene families.

Complex carbohydrates of plants are the main substrates for white-rot fungi. Carbohydrate-active enzymes (CAZymes) play a critical role in complex carbohydrates metabolism as well as many other important physiological processes, such as development and stress response (Xie et al., 2018; Huang et al., 2020). The CAZymes of the other 22 fungal species were analyzed against the dbCAN2 database with the same parameters. A total of 541 genes accounting for 577 CAZymes terms were identified in the L. decastes genome, including 253 GHs, 77 GTs, 24 PLs, 40 CEs, 154 AAs, and 29 CBMs (Figure 4A; Supplementary Table 5).

Lyophyllum decastes was a wood-rooting white-rot fungus. The media for L. decastes production conpose mainly lignin-rich media. It suggested that L. decastes has powerful lignin modification potential. AA families in the CAZymes database are mainly involved in depolymerization of non-carbohydrate structural components, lignin. Proteins in the AA families were mainly distributed in AA3, AA9, AA1, AA7, AA5, and AA2 categories. Laccases (AA1 family) and class II peroxidases (AA2 family) are important for the degradation of lignin in lignocellulose. A total of 25 laccases in the AA1 category (23 AA1_1, 1 AA1, and 1 AA1_2 category) and 7 class II peroxidases were annotated in the AA2 category in L. decastes genome (Supplementary Table 5). The number of laccases is the highest among all the analyzed fungi, which indicated that lignin degradation by L. decastes depends mainly on the addition of AA1 subfamilies. The results confirmed the lignin degradation potential of this mushroom, which is consistent with the wood-rich media used in L. decastes production.

Interestingly, Termitomyces species showed a close evolutionary relationship with L. decastes. CAZymes analysis revealed that only 87 AAs (56.5% of those annotated from L. decastes) were identified in Termitomyces sp., including 16 from the AA1_1 category and 3 from the AA2 category (Figure 4B). Different from species of Lyophyllaceae, Termitomyces are symbiotic fungi, which depend on fungus-growing termites. Termites use their fecal pellets from plant materials as the growing media to cultivate Themitomyces mycelia (Aanen et al., 2007). Termites feed on lignocellulose, remove 74%–99% of the cellulose and 65%–87% of the hemicellulose components, and defecate lignin-rich feces (Brune, 2014). Also the media of Termitomyces sp. is mainly lignin-riched feces, the accessibility of cellulose is higher than those of raw lignocellulose materials. Therefore, the lignin-degrading enzymes were lost during the evolution.

In addition to AAs, 253 proteins from GH family were identified in the genome L. decastes. There proteins were subclassified into 55 GH families and they were mainly distributed in GH5, GH18, GH3, and GH43 families. Proteins in GH families were cellulose and hemicellulose degradation related enzymes, which suggested that L. decastes could use cellulose and hemicellulose as the substrate for mycelial growth.

The mating loci of L. decastes were identified using Blastp tools against mating-related proteins in relative species such as H. marmoreus and P. giganteus. Five HD genes, including 2 HD1 genes and 3 HD2 genes, were identified on chr01 in L. decastes strain LRG-d1-1 (Figure 5A). The five HD genes are grouped into two pairs of homeodomain transcription factor genes, and the organization of these genes is similar to the type1 mating A locus in H. marmoreus (Wang et al., 2021a). The mating A locus of strain LRG-d1-2 and strain LRG-d1-5 has four HD genes grouped into two pairs of HD transcription factor genes. The organization of these genes is different from all kinds of mating A loci in H. marmoreus reported by Wang et al. (2021a). The mitochondrial intermediate peptidase (Mip) and beta flanking gene (Bfg) were located in the two ends of the mating A locus of L. decastes. The mating B locus was identified on chr10 in L. decastes strain LRG-d1-1 (Figure 5B). The mating B loci of the three L. decastes strains all have five STEs. The organization of these genes in the three strains is similar, while different from mating B loci reported in H. marmoreus (Wang et al., 2021).

According to the nucleotide sequence of mating loci in strain LRG-d1-1 and LRG-d1-5, four pairs of primers (Supplementary Table 2) were designed for the PCR-based verification of L. decastes mating types. Applicability of the primers was tested in the monokaryotic mycelial isolates raised from the basidiospores of L. decastes LRG-d1 and the dikaryotic L. decastes strains. Among 8 monokaryotic isolates, all strains yielded two DNA bands either from mating A or from mating B (Supplementary Figure 3). According to PCR results, the strain LRG-d1-1 is A1B1 mating type, while the LRG-d1-2 and LRG-d1-5 are A2B2 mating type, in agreement with the genome information. The three dikaryotic isolates showed four DNA bands from A1, A2, B1, and B2 mating types (Supplementary Figure 3). The results confirmed that the four primers could precisely distinguish the two different mating types of L. decastes.

To further validate the effectiveness of the primers in cross-breeding, mating experiments based on PCR results were performed. The monokaryotic isolates with different A mating types and different B mating types were placed on one PDA plate at a close distance. After 2 weeks of incubation, mycelial samples from the growing edge on the perpendicular bisector between two inoculation blocks. The mating type of the samples was measured using PCR with the mating type-specific primers and the micromorphology of the samples was checked using a microscope. Our results confirmed that the monokaryotic strains with compatible mating types could form fertile dikaryotic mycelia with clamp connections (Supplementary Figure 3). The results confirmed that L. decastes is a tetrapolar heterothallism mating type species. Identification of the exact mating type of a monokaryotic strain can reduce the cross-breeding time. Therefore, the results presented in this study contribute to the L. decastes breeding industry through accelerating of the cross-breeding process.

As a wood-rotting fungus, farmers normally use lignin-rich sawdust from broad-leaved trees and cotton hull as carbon sources and use bran as the nitrogen source for L. decastes fruiting body production. Efficient lignocellulose utilization is one of the key factors to increase mushroom yield. However, the ability of L. decastes to degrade lignocellulose is still not clear. To investigate the molecular mechanism underlining lignocellulose degradation by L. decastes, the secretome of L. decastes in PDB media (PDB group), PDB + bran media (Bran group), PDB + cotton hull media (Cott group), and PDB + Sawdust media (Sawd group) were analyzed (Figure 6A) using label-free proteomic analysis.

A total of 1,071 proteins were identified in 12 samples from L. decastes supernatant of 4 groups (three biological replicates for each group; Supplementary Table 6), and 415, 752, 677, and 782 proteins were identified in the PDB, Bran, Cott, and Sawd groups, respectively (Figure 6B). Among identified proteins, only 23.6% (253) proteins were identified in all groups. Interestingly, 18.0% (193) proteins were identified in three lignin-rich groups but not in the PDB group. It is worth noting that the protein number identified in lignin-rich media were much higher than those in the PDB group. However, the number of secreted proteins or CAZymes are similar between different groups (Figure 6E). A similar result was also observed in P. ostreatus that straw and poplar revealed a strong and common effect of lignocellulose on the secreted proteins (Fernandez-Fueyo et al., 2016). The result indicated that all three lignin-rich substrates could induce the expression of a more complex secretome than PDB media. PCA results revealed that samples from the same group clustered while samples from different groups were divided into four groups by the first and second components (Figure 6C). Pearson correlation coefficients analysis revealed similar results, that samples from the same group have high correlation values while samples from different groups have lower correlation values (Figure 6D). The results suggested that the protein expression levels induced by different substrates were different despite similar protein numbers identified in the three lignin-rich groups.

Four media used in this study were all derived from PDB media, which contain 20 g/L (111 mM) glucose. High glucose concentration could inhibit cellulose hydrolysis activity and the utilization of hemicellulose (Ohmine et al., 1983; Nijland et al., 2019). Therefore, a high concentration of glucose might inhibit the expression of cellulose and hemicellulose degradation enzymes by so-called feedback inhibition. As expected, the expressions of cellulose and hemicellulose degradation enzymes in L. decastes were not significantly induced by the lignin-rich substrates. Twenty-one cellulose degradation enzymes from GH families were identified through proteomic analysis. However, the total abundance in the lignin-rich groups is not induced compared with the PDB group. Xylan hemicelluloses are considered the second most abundant class of polysaccharides after cellulose. Three xylosidases and one xylanase were identified through proteomic analysis (Supplementary Table 7), and three were only identified in lignin-rich groups. Alpha-xylosidase, MDBLdec1_11857, has the highest abundance among the four proteins, and it was identified in all three lignin-rich goups but PDB group. The results indicated that xylan in lignocellulose substrates might induce the expression of xylan degradation enzymes in lignin-rich groups. Even so, the abundance of these enzymes is quite low.

Laccases (AA1 CAZymes family) and class II peroxidases (AA2 CAZymes family) are two major classes of enzymes involved in the modification of lignin. Three class II peroxidases and 17 laccases were identified from four proteome groups. The abundances of these genes were shown with a heatmap plot in Figure 7. Obviously, the abundances of two laccases (MDBLdec1_07135, MDBLdec1_09727) in the lignin-rich groups are significantly higher than those in the PDB group and much higher than those of other identified laccases. Furthermore, the abundances of these two laccases are among the highest-three abundance proteins in lignin-rich groups (Supplementary Table 7). Similar results have also been observed in other white-rot fungi. Fernández-Fueyo reported that the abundance of laccase increased from 1% of the total protein abundances in non-lignin media to 21% in the lignin-rich media (Fernandez-Fueyo et al., 2016). Therefore, MDBLdec1_07135 and MDBLdec1_09727 are considered to play essential roles in the lignin degradation in L. decastes. MDBLdec1_07135 and MDBLdec1_09727 show the highest sequence identity with 66,615 and 123,288 proteins in PC9, respectively. However, the two proteins were not identified in the secretome in P. ostreatus PC9 (Fernandez-Fueyo et al., 2016), suggesting different regulation pathways between the two species. Besides, some other laccases, such as MDBLdec1_09748, MDBLdec1_08907, and MDBLdec1_07134, also show relatively high expression abundance in four groups; however, these laccases were not induced in lignin-rich media. Starch usually co-exists with lignocellulose in nature, therefore, the expression of these laccases might be induced by the starch or other polymer in potato extract.

Heatmap of the AA2 family proteins revealed that two peroxidases (MDBLdec1_05507, MDBLdec1_11977) were only expressed in lignin-rich groups. The abundance of MDBLdec1_02349 in the Cott group is significantly higher than that in the PDB group. The results indicated that despite the different structures of the lignocellulose substrates of the three lignin-rich groups the expression patterns of laccases and class II peroxidases in the three groups are similar. The three substrates might produce the same inducer for the induction expression of these enzymes. Peroxidase catalyzing reaction requires H₂O₂ as the co-substrate for lignin degradation. White rot fungi secreted many partner enzymes that co-expressed with peroxidases, such as glucose-methanol-choline oxidoreductases from the AA3 CAZymes family, glyoxal oxidases (AA5 family) and oligosaccharide oxidases (AA7 family). In our proteomic analysis, 23, 6, and 7 proteins from AA3, AA5, and AA7 families were identified from four groups (Figure 7), respectively. These enzymes oxidize a variety of carbohydrates, alcohols, and aldehydes derived from lignocellulose. The results indicated that three lignocellulose substrates, sawdust, cotton hull, and bran, were degraded to different products, and the expression of these peroxidase partner enzymes may be induced by corresponding substrates. H₂O₂ could not only serve as a co-substrate it could also inhibit the lignin-active peroxidase activity. Therefore, white rot fungi produced the H₂O₂-scavenging enzyme, catalase, to regulate the high concentration of H₂O₂. Two catalases, MDBLdec1_06794 and MDBLdec1_13173 were identified in proteomic analysis. Interesting, MDBLdec1_13173 abundances in Bran and Sawd groups were significantly higher than that in PDB and Cott groups and MDBLdec1_06794 was only expressed in Bran and Sawd groups (Figure 7). The results indicated that different substrates induced different peroxidase expression patterns, which further influence the expression of partner enzymes such as H₂O₂-generating and H₂O₂-scavenging partner enzymes. Besides, catalases expression are usually associated with stress response in vivo. Therefore, the extracellular expression of catalases might also influenced by the stress response in mycelia.

Laccases and class II peroxidases transform lignin into reactive quinones and phenoxy radicals in vitro, which leads to the formation of quinone and (re)polymerized lignin. To efficiently depolymerize lignin, partner enzymes in the white-rot fungi secretome are required to deactivate laccase- and peroxidase-generated quinones and phenol radicals (Monclaro et al., 2022). These enzymes include dehydrogenases from the AA3 CAZymes family and quinone reductases from the AA6 CAZymes family. Here, we identified an AA6 family quinone reductase (MDBLdec1_12289) in proteomic analysis. Compared with the abundance of MDBLdec1_12289 in the PDB group, the abundances in Bran, Cott, and Sawd groups increased by 12.4-fold, 27.7-fold, and 12.4-fold, respectively (Figure 7). It was reported that AA6 quinone reductase could increase the lignin depolymerization by Lip in vitro (Majeke et al., 2021). The results suggested that MDBLdec1_12289 played important role in lignin decomposition in L. decastes.

Laccases generally oxidize a wide range of substrates, such as lignin; typically transform phenols and aromatic amines into phenoxy radicals. These unstable chemical products commonly start domino reactions to form phenols or quinones (Arregui et al., 2019; Janusz et al., 2020). Quinones can be used by quinone reductases, laccases, and peroxidases in redox cycle reactions to activate oxygen which could further produce hydroxyl radicals. Hydroxyl radicals are among the strongest oxidants in white-rot fungi cultures and can initiate the attack on lignocellulose, which leads to the depolymerization of lignin (Gasser et al., 2012). Cuillen et al. reported the existence of quinone redox cycling in white-rot fungi Pleurotus eryngii. Therefore, the results suggested that the two laccases (MDBLdec1_07135 and MDBLdec1_09727) and one quinone reductase (MDBLdec1_12289) play an essential role and have a synergistic effect on lignin decomposition in L. decastes LRG-d1 (Guillen et al., 1997). However, further work is needed to characterize the role of laccase and quinone reductases in lignin decomposition.