Comparative Proteomic Analysis of Flammulina filiformis Reveals Substrate-Specific Enzymatic Strategies for Lignocellulose Degradation

Abstract

Flammulina filiformis, one of the most delicious and commercially important mushrooms, demonstrates remarkable adaptability to diverse agricultural wastes. However, it is unclear how different substrates affect the degradation of lignocellulosic biomass and the production of lignocellulolytic enzymes in F. filiformis. In this study, label-free comparative proteomic analysis of F. filiformis cultivated on sugarcane bagasse, cotton seed shells, corn cobs, and glucose substrates was conducted to identify degradation mechanism across various substrates. Label-free quantitative proteomics identified 1104 proteins. Gene Ontology and Kyoto Encyclopedia of Genes and Genomes pathway analysis of protein expression differences were predominantly enriched in energy metabolism and carbohydrate metabolic pathways. Detailed characterization of carbohydrate-active enzymes among the identified proteins revealed glucanase (GH7, A0A067NSK0) as the key enzyme. F. filiformis secreted higher levels of cellulases and hemicellulases on sugarcane bagasse substrate. In the cotton seed shells substrate, multiple cellulases functioned collaboratively, while in the corn cobs substrate, glucanase predominated among the cellulases. These findings reveal the enzymatic strategies and metabolic flexibility of F. filiformis in lignocellulose utilization, providing novel insights for metabolic engineering applications in biotechnology. The study establishes a theoretical foundation for optimizing biomass conversion and developing innovative substrates using targeted enzyme systems.

1. Introduction

F. filiformis (commonly known as winter mushroom), a basidiomycete fungus, belongs to the class Hymenomycetes, order Agaricales, family Tricholomataceae, and genus Flammulina [1]. This edible mushroom contains abundant bioactive compounds, including immunomodulatory proteins, fungal polysaccharides, ergothioneine, flavonoids, terpenoids, dietary fiber, vitamins, and essential trace elements. These components confer multiple pharmacological benefits, such as immune regulation, anti-tumor activity, and cholesterol-lowering effects [2,3]. Currently, F. filiformis has gained increasing popularity as a nutritious food source due to its low-calorie content, high dietary fiber, and a rich nutrient profile. Its demand has been steadily growing in consumer markets, particularly across Asian and European regions [4,5].

Currently, the primary lignocellulosic substrates for F. filiformis cultivation in China are corn cobs and cotton seed shells. Corn cobs serve as the main carbon source for F. filiformis cultivation [6], with cellulose constituting approximately 30% of their composition [7]. Upon decomposition, these components provide essential nutrients for fungal growth and development [8]. Cotton seed shells, widely utilized in edible mushroom cultivation due to their balanced carbon-to-nitrogen ratio and favorable physical properties, are rich in crude protein, crude fat, and crude fiber [9]. Sugarcane bagasse primarily contains cellulose, accounting for about 33%~50%, hemicellulose, and lignin [10]. As a major byproduct of the sugar industry, sugarcane bagasse is predominantly used in biomass combustion, fertilizer production, ethanol fermentation, and biofeed manufacturing, demonstrating abundant availability and untapped potential despite suboptimal utilization rates. Numerous studies have reported its application in mushroom cultivation [11,12], where it effectively enhances yield and biological efficiency as a carbon source [13]. Consequently, lignocellulose in substrates represents a critical nutrient for F. filiformis growth, and the fungal capacity to degrade lignocellulose significantly is one of the key factors affecting their cultivation performance.

Lignocellulose, the most abundant biomass resource on Earth, is composed of lignin, cellulose, hemicellulose, and minor components, including pectin, proteins, and ash [14]. Cellulose, constituting 40–50% of lignocellulose, is the predominant component in lignocellulose. It is a linear macromolecule formed by tens of thousands of glucose residues linked via β-1,4-glycosidic bonds [15,16]. F. filiformis, a typical white-rot fungus, degrades lignocellulose by secreting lignocellulolytic enzymes, thereby providing energy for substance synthesis, transport, and accumulation during mycelial growth [17]. Enzymes involved in the synthesis and degradation of glycoconjugates, oligosaccharides, and polysaccharides are collectively termed carbohydrate-active enzymes (CAZymes). CAZymes are classified into six categories: glycosyl transferases (GTs), carbohydrate esterases (CEs), glycoside hydrolases (GHs), polysaccharide lyases (PLs), auxiliary activities (AAs), and carbohydrate-binding modules (CBMs) [18]. Lignocellulolytic enzymes, belonging to the CAZyme family, play an important role in the growth of white-rot fungi [19]. Based on functionality, lignocellulolytic enzymes are categorized into three groups: ligninase, cellulase, and hemicellulase.

Cellulases refer to a multicomponent enzyme system capable of hydrolyzing β-D-glycosidic bonds in cellulose to yield glucose. These enzymes are composed of endoglucanase (EC 3.2.1.4), exoglucanase (EC 3.2.1.91), and β-glucosidase (EC 3.2.1.21), which collectively function as GHs to comprehensively degrade cellulose into glucose [20].

Hemicellulose is composed of heteropolysaccharides such as xylan, arabinan, galactan, and mannan, exhibiting an amorphous and randomly coiled spatial conformation with lower degrees of polymerization and molecular weights compared to cellulose. Hemicellulases include endo-1,4-β-xylanase (EC 3.2.1.8), β-1,4-xylosidase (EC 3.2.1.37), endo-1,4-β-galactosidase (EC 3.2.1.89), β-galactosidase (EC 3.2.1.23), β-mannosidase (EC 3.2.1.25), endo-α-1,5-arabinanase, and α-N-arabinofuranosidase (EC 3.2.1.55) [21].

The enzymatic systems responsible for lignin degradation in fungi primarily include laccase (EC 1.10.3.2), manganese peroxidase (EC 1.11.1.13), lignin peroxidase (EC 1.11.1.14), and versatile peroxidase (EC 1.11.1.16) [22].

The decomposition, utilization, and conversion of lignocellulose by fungi represent a highly complex process requiring the coordinated action of multiple extracellular enzymes [23]. Studies have reported that basidiomycetes, exemplified by Ganoderma lucidum, induce distinct combinations of lignocellulolytic enzymes in response to variations in lignin, cellulose, and hemicellulose content and structural composition within growth media, and there are expression differences among the genes of these enzymes [21], which are ultimately reflected in the secretory proteome. Proteomics research can elucidate condition-specific protein expression profiles, revealing decomposition strategies of fungi on different substrates through functional and relational analyses of extracellular proteins. Research on the degradation enzyme activities of cellulose, hemicellulose, and lignin in F. filiformis under conditions of different carbon sources and nitrogen sources using corn cobs and cotton seed shells revealed that under simple carbon source conditions, the activities of carboxymethyl cellulase (CMCase) and xylanase were lower than those in complex carbon source media. In the complete medium, the activities of these two enzymes were lower than in carbon- and nitrogen-deficient media. Moreover, laccase activity was lower in media without simple nitrogen sources compared to the complete medium and media without glucose, indicating that the laccase activity is lower in media with complex carbon and nitrogen sources than in those with simple carbon and nitrogen sources [24].

In a complementary study, Pleurotus ostreatus mobilized substrate-specific antioxidant systems and carbon metabolic pathways to produce an enhanced repertoire of ligninolytic enzymes when cultivated on lignocellulose-rich biomass. Comparative analyses revealed that P. ostreatus grown on sawdust exhibited elevated expression of ligninolytic enzymes and CAZymes, whereas the highest transcriptional upregulation of CAZymes was detected in cotton seed shells and corn cobs substrates. These findings demonstrate that P. ostreatus adapts to heterogeneous lignocellulosic matrices through differential protein expression modulation, thereby optimizing its growth and developmental efficiency under varying substrate conditions [25]. Proteomic analysis of the secretory proteins from Pleurotus eryngii mycelia cultured in liquid media containing sawdust, bagasse, peanut shells, and glucose revealed that the most prevalent CAZymes involved in lignocellulose degradation across all three substrates were laccase (Lac), manganese peroxidase (MnP), aryl alcohol oxidase (AaO), and copper radical oxidase (CRO). Among these, Lac2 was predominantly implicated in the degradation of sawdust, peanut shells, and sugarcane bagasse, while Mnp3 played a central role in lignin depolymerization within peanut shell substrates. Functional characterization further demonstrated that AaO and Lac4 primarily contributed to oxidative stress responses during glucose catabolism, serving critical protective roles in mitigating oxidative damage to mycelia [26]. In-depth investigation of the secretory proteins in lignocellulose-degrading fungi holds significant scientific and practical value for deciphering their metabolic pathways and molecular mechanisms underlying lignin biodegradation.

In this study, F. filiformis was cultivated on four distinct carbon sources (glucose, cotton seed shells, corn cobs, and sugarcane bagasse), and label-free quantitative proteomics was employed to characterize its secretory protein profiles and lignocellulolytic enzyme systems. The analysis revealed carbon source-dependent expression patterns of key enzymatic components, facilitating the elucidation of molecular strategies for lignocellulose degradation and utilization in F. filiformis. These findings provide critical insights into fungal adaptation mechanisms to heterogeneous substrates and offer valuable references for optimizing lignocellulosic biomass-based cultivation systems.

2. Materials and Methods

2.1. Fungal Strain

The experimental strain F. filiformis WYS-20 was provided by the Institute of Agricultural Resources and Environment, Shandong Academy of Agricultural Sciences

2.2. Reagents and Apparatus

Potato dextrose agar (PDA) powder (Becton, New York, NJ, USA), glucose (Becton, New York, NJ, USA), sugarcane bagasse, cotton seed shells, and corn cobs were used as substrates. These substrates were obtained through mechanical milling of particles, resulting in particles with a size <5 mm. The three substrates were obtained from Lvyuan Yongle Agricultural Technology Development Co., Ltd. (Beijing, China).

2.3. Culture Conditions

Corn cobs, sugarcane bagasse, and cotton seed shells are used as additives in liquid substrates. These substrates are obtained through mechanical grinding into particles with a size of less than 5 mm. Before addition, they were sterilized at 126 °C for 30 min. For the liquid medium preparation, the base medium per liter contained 20 g glucose, 2 g peptone (AOBOX, Beijing, China), 1 g KH₂PO₄ (Solarbio, Beijing, China), and 0.5 g MgSO₄ (Xi Long Scientific, Guangzhou, China). In the other three experimental groups, glucose was replaced with equivalent quantities of distinct agricultural waste materials (sugarcane bagasse, cotton seed shells, and corn cobs).

Mycelial plugs (5 mm diameter) of the F. filiformis strain WYS-20 were inoculated onto PDA plates and incubated at 21 °C for 7 days. For each treatment group (the glucose control group and three types of agricultural waste materials groups), three independent biological replicates were established, with each replicate utilizing mycelial plugs originating from discrete sources and undergoing separate cultivation. Under aseptic conditions, mycelia with agar medium from each plate were transferred to a homogenizer, blended with 100 mL of liquid medium using intermittent pulses for 30 s, and separately cultured in 250 mL Erlenmeyer flasks. The cultures were maintained in darkness at 180 rpm for 14 days. Post-cultivation, the fermented broth was centrifuged at 13,000× g for 10 min, and the supernatant was clarified through 0.45 μm membrane filters. For protein quantification and proteomic analysis, 50 mL of filtrate was collected from each sample.

2.4. Secreted Protein Preparation

Referring to the study of Pleurotus eryngii [26], before conducting the proteomic analysis, we performed biomass normalization for all samples. The specific steps are as follows: First, we quantified the biomass by measuring the dry mycelial weight. Then, we adjusted the protein extraction amount for each sample according to the dry weight ratio to ensure that all samples had the same biomass basis for proteomic analysis. This normalization step helps eliminate potential biases arising from differences in biomass, thereby enhancing the comparability of the proteomic data. After protein quantification, 60 μg of protein solution was placed in a centrifuge tube, followed by the addition of 5 μL of DTT (Genview, Naples, FL, USA) (1 mol·L⁻¹) solution and incubation at 37 °C for 1 h. Subsequently, 20 μL of IAA (Sigma-Aldrich, Oakville, ON, Canada) (1 mol·L⁻¹) solution was added, mixed adequately, and allowed to react for 1 h at room temperature away from light. All samples were aspirated into an ultrafiltration tube and centrifuged, and the collection fluid was discarded. This was followed by the incorporation of 100 μL of UA (8 M urea, 100 mM Tris-HCl, pH = 8.0) (Sigma-Aldrich) into an ultrafiltration tube, and repeating centrifugation twice. Afterward, 100 μL of NH₄HCO₃ (Sigma-Aldrich) (0.05 mol·L⁻¹) was added and centrifuged, and the collection solution was discarded, and the process was repeated three times. A new collection tube was used for tryptic digestion. Trypsin was added at a 50:1 ratio of protein to the enzyme, and digestion proceeded at 37 °C for 12–16 h.

2.5. LC-MS/MS Data Acquisition

Soluble extracellular proteins were subjected to mass spectrometry analysis following the methodology established for Pleurotus ostreatus [27]. The samples after enzymatic hydrolysis were pressure-loaded onto a fused silica capillary chromatography column packed with 3 μm Dionex C18 material (RP; Phenomenex, Torans, CA, USA). The reversed-phase column segment (pore size 100 Å) has a length of 15 cm. The column was washed using buffer A (water containing 0.1% formic acid) and buffer B (acetonitrile (ACN) containing 0.1% formic acid). After desalting, a 5 mm long, 300 µm inner diameter C18 capture tip was coupled in series with an Agilent 1100 quaternary high-performance liquid chromatography (HPLC) system and analyzed using a 12-step separation method. The first step of the gradient elution program consisted of a 5 min increase from 0% buffer B to 2% buffer B, followed by a 45 min gradient to 40% buffer B. Next, buffer B was increased from 40% to 80% over 3 min and maintained at 80% for 10 min. After that, buffer B was decreased from 80% to 2% over 2 min. Approximately 100 μg of the tryptic peptide mixture was then loaded onto the chromatography column and separated using a linear gradient at a flow rate of 0.5 μL/min. When the peptides were eluted from the microcapillary chromatography column, they were directly ionized by electrospray at a source temperature of 180 °C into the micrOTOF-Q II mass spectrometer (Bruker Scientific, Billerica, MA, USA). The mass spectrometer operated in MS/MS (auto) mode. Full scan mass spectra (Survey MS scans) were collected on the TOF-Q II (Bruker Scientific, Billerica, MA, USA), with the resolution set to 20,000. Following each full scan (mass-to-charge ratio range m/z 50-2500), five data-dependent tandem mass spectrometry (MS/MS) scans were performed, with a normalized scan rate of 2 Hz.

2.6. Sequence Database Search and Data Analysis

Data analysis was performed using Proteome Discoverer 2.1 software (Thermo Fisher Scientific, Waltham, MA, USA, version 2.1). Peptide identification was conducted using the SEQUEST search engine, searching against a human proteome database containing reviewed sequences downloaded from UniProt (** entries). The decoy database for the search was generated using the revert function. The parameters set for protein identification were as follows: peptide mass tolerance = ±15 ppm, MS/MS tolerance = 0.02 Da, enzyme = trypsin, maximum missed cleavages allowed = 2, fixed modification: carbamidomethylation (C), variable modification: oxidation (M), and database search mode = decoy. The false discovery rate (FDR) threshold at both peptide and protein levels was set at 0.01.

Proteins showing upregulation or downregulation in both replicates, with a relative quantitative p-value < 0.05 and a fold change ≥ 1.5, were selected as differentially expressed proteins (DEPs). Experimental data from three technical replicates performed in triplicate were analyzed.

2.7. Statistical Analysis

Statistical analysis of the database search results obtained from MaxQuant (version 2.0.1.0) was performed using Perseus software (version 1.3.0.4). Differentially expressed proteins were functionally annotated and subjected to pathway analysis via the Gene Ontology (GO) classification system (http://geneontology.org/ (accessed on 2 August 2021)) and the Kyoto Encyclopedia of Genes and Genomes (KEGG) online bioinformatics tool (https://www.kegg.jp/ (accessed on 2 August 2021)). GO enrichment analysis was conducted using Fisher’s exact test to identify statistically significant functional categories.

3. Results

3.1. Differential Protein Expression Profiles

The protein expression profiles of F. filiformis cultivated on four different substrates were analyzed using LC-MS/MS proteomics. After culturing on the four substrates, proteins were extracted, reduced, alkylated, and subjected to enzymatic digestion, followed by high-resolution mass spectrometry for peptide data acquisition.

Database searching using Proteome Discover (version 2.4) identified 984, 922, 745, and 681 proteins in samples from the sugarcane bagasse, cotton seed shells, corn cobs, and glucose substrates, respectively. A total of 1104 differentially expressed proteins were identified across all substrates, as shown in Figure 1. Venn diagram analysis reveals that 809 proteins were commonly expressed across all four substrates, with the highest number of proteins identified in the sugarcane bagasse. The full protein list (Table S1) is deposited in Supplementary Materials.

Proteins identified in this study were analyzed using the dbCAN and CAZymes databases to reveal the distribution of glycoside hydrolase families (GHs) in F. filiformis cultivated on four substrates. Analysis of the identified proteins shows the distribution of proteins in secreted protease families across the four substrates, and the results are presented in

Table 1. Annotation results of carbohydrate-active enzymes in secretory proteins of Flammulina filiformis cultivated on four substrates.

Protein ID	CAZy Family	Protein Description	Signalp	Protein Abundance
				Glucose	Sugarcane Bagasse	Corn Cobs	Cotton Seed Shells
A0A067N481	GH5	Glycoside hydrolase family 5 protein	N
A0A067N5C6	GT2	Chitin synthase	N	1,674,382.92233333	1,553,722.012	761,452.014833333	1,626,613.11766667
A0A067N7N9	AA1	Fet3 ferroxidase	Y	1,923,473.892	1,569,898.35733333	814,216.989533333	1,917,835.20633333
A0A067N9N5	GT35	Alpha-1,4 glucan phosphorylase	N	5,045,299.339	2,661,980.72466667	3,300,366.021	2,613,296.99
A0A067N9V0	GH3	Glycoside hydrolase family 3 protein	N	73,966,935.1533333	65,251,685.5266667	42,499,127.7966667	125,931,711.033333
A0A067NB35	GH35	Glycoside hydrolase family 35 protein	N	44,803,576.2366667	54,079,091.2	40,553,862.6666667	62,096,653.43
A0A067NCX6	GH35	Beta-galactosidase	N	26,270,162.46	12,170,355.1233333	9,266,884.40666667	4,327,699.927
A0A067NDH9	GH16	Glycoside hydrolase family 16 protein	N	705,834.522833333	589,586.945666667	724,761.576	1,811,456.84333333
A0A067NE50	PL26	Autophagy-related protein 27	Y	37,554,727.2766667	27,057,562.07	13,562,714.79	13,853,933.0233333
A0A067NG33	GT3	Glycogen [starch] synthase	N	8,748,214.16666667	1,328,355.905	2,525,576.068	1,244,528.46533333
A0A067NG43	GH179	GFO_IDH_MocA domain-containing protein	N	687,454.406266667	1,619,932.57866667	807,516.327666667	1,428,833.614
A0A067NG45	GT69	Glycosyltransferase family 69 protein	Y	4,409,595.69266667	4,622,098.876	2,747,698.281	2,672,916.48933333
A0A067NHG4	GH51	Non-reducing end alpha-L-arabinofuranosidase	N	1,153,395.48766667	3,097,140.86366667	3,939,891.408	10,696,011.0986667
A0A067NHT6	GH5	Glycoside hydrolase family 5 protein	Y	43,335.89372	761,726.745733333	219,575.423666667	43,335.89372
A0A067NHZ2	AA8	GMC_OxRdtase_N domain-containing protein	N	545,214.741766667	91,463,449.0566667	6,676,565.561	2,890,166.06066667
A0A067NI47	GH30	Glycoside hydrolase family 30 protein	N	2,387,567.25	215,776.937866667	36,963.9312	36,963.9312
A0A067NIA1	CE15	Carbohydrate esterase family 15 protein	Y	339,189.8876	115,408,491.666667	41,968,924.2733333	8,447,431.35666667
A0A067NJ89	GH5	Glycoside hydrolase family 5 protein	N	284,593.773466667	326,019.267	115,600.020666667	21,217.80416
A0A067NJZ2	GH115	Glycoside hydrolase family 115 protein	Y	1,930,693.30433333	4,595,319.543	1,213,666.526	369,670.60656
A0A067NKG1	GH63	Glyco_hydro_63 domain-containing protein	N	2,747,527.69166667	983,334.406933333	1,246,596.6286	1,117,502.767
A0A067NKR6	GH3	Glycoside hydrolase family 3 protein	N	1,172,579.32166667	4,990,872.88133333	2,641,664.84333333	7,850,058.13933333
A0A067NL12	CE4	Chitin deacetylase	Y	2,532,754.30033333	924,863.084666667	1,155,317.05666667	2,255,575.432
A0A067NL90	AA3	GMC_OxRdtase_N domain-containing protein	Y	882,786.094366667	2,610,349.02066667	808,885.0888	3,630,119.53566667
A0A067NLS2	GH38	Alpha-mannosidase	N	7,706,159.83633333	12,219,068.8266667	1,657,370.85133333	1,078,749.07483333
A0A067NNJ9	GH13	1,4-alpha-glucan-branching enzyme	N	1,242,357.576	29,290.3292333333	302,425.2643	109,230.33996
A0A067NNT3	GT15	Glycosyltransferase family 15 protein	N	1,208,833.28266667	1,664,963.70766667	552,614.189366667	210,154.392233333
A0A067NP57	GH74	Glycoside hydrolase family 74 protein	Y	388,005.1119	47,425,233.0866667	7,718,332.32966667	22,058,456.8833333
A0A067NPC3	GH20	Beta-hexosaminidase	Y	40,254,045.4666667	14,772,757.8533333	9,881,569.315	24,482,395.38
A0A067NQN1	GH74	Glycoside hydrolase family 74 protein	Y	463,429.509266667	58,716,335.94	10,340,246.84	25,877,955.2
A0A067NRB1	GH3	Beta-glucosidase	N	82,215.74544	2,068,114.20433333	1,022,417.6928	1,592,823.36066667
A0A067NRS6	AA4	FAD-binding PCMH-type domain-containing protein	N	12,488,633.6083333	9,745,718.80066667	5,172,592.48033333	40,775,298.46
A0A067NSK0	GH7	Glucanase	Y	11,217,736.1	1,268,879,946	230,125,783.033333	923,739,948.666667
A0A067NTR2	GH13	Glycoside hydrolase family 13 protein	Y	285,318.224566667	502,909.6734	212,830.326166667	186,036.48156
A0A067NTS1	GH72	1,3-beta-glucanosyltransferase	N	54,130,175.98	4,963,599.49233333	11,825,230.4266667	105,578,133.73
A0A067NU72	GH47	alpha-1,2-Mannosidase	N	1,769,389.19833333	280,684.350966667	372,926.358866667	49,006.6803
A0A067NUA1	GH3	Glycoside hydrolase family 3 protein	N	795,122.5487	127,320.676066667	929,881.6146	517,719.2041
A0A067NUJ5	GT20	Glycosyltransferase family 20 protein	N	4,084,710.286	810,559.760733333	548,366.213233333	561,918.323533333
A0A067NUM9	GH43	Glycoside hydrolase family 43 protein	N	1,511,309.05833333	59,552,658.17	56,268,782.9433333	76,230,697.6633333
A0A067NVF4	GH47	alpha-1,2-Mannosidase	Y	328,118.7679	632,440.375066667	251,327.522866667	46,763.0276
A0A067NVK8	GH179	GFO_IDH_MocA domain-containing protein	N	972,190.939133333	796,142.437633333	834,063.037033333	2,027,235.743
A0A067NW26	GH27	Alpha-galactosidase	N	231,569.9718	2,884,487.07166667	35,724.60152	35,724.60152
A0A067NXJ5	GT69	Glycosyltransferase family 69 protein	N	579,444.1575	772,410.943	1,040,558.4363	615,105.0206
A0A067NXX6	GH5	Glycoside hydrolase family 5 protein	N	142,572.07057	425,257.856066667	308,028.0275	537,522.797633333
A0A067NYC9	AA9	Glycoside hydrolase family 61 protein	N	32,413.33185	191,864.378366667	98,566.1137166667	516,223.5153
A0A067NYD8	GT39	Dolichyl-phosphate-mannose--protein mannosyltransferase	N	184,289.4021	256,503.235966667	45,563.4125	45,563.4125
A0A067NYI6	AA9	Glycoside hydrolase family 61 protein	Y	111,182.064	1,637,476.95666667	595,988.317233333	111,182.064
A0A067P039	GH3	Beta-glucosidase	N	496,367.998333333	5,285,172.74033333	2,872,587.55566667	1,756,140.254
A0A067P0D5	GT2	Chitin synthase	N	115,901.747933333	156,000.033433333	20,483.6242	20,483.6242
A0A067P120	GH2	Glycoside hydrolase family 2 protein	N	616,712.845333333	687,912.198833333	213,204.624633333	126,479.038473333
A0A067P222	GH3	Beta-glucosidase	N	463,925.3207	12,174,613.98	5,160,192.30733333	3,392,101.14166667
A0A067P2G6	GH74	Glycoside hydrolase family 74 protein	N	1,029,203.9254	92,302,563.21	12,674,182.64	29,404,276.5166667
A0A067P3G0	GH43	Glycoside hydrolase family 43 protein	Y	2,249,040.07066667	1,483,016.085	334,123.3293	61,306.25716
A0A067P3W7	GH10	Beta-xylanase	Y	1,473,446.39433333	196,961,002.933333	25,016,172.1333333	24,481,648.0066667
A0A067P3Z6	GH5	Glycoside hydrolase family 5 protein	Y	22,239,740.3866667	12,050,158.5566667	14,542,128.2233333	26,057,009.0033333
A0A067P4G8	PL8	Polysaccharide lyase family 8 protein	Y	11,472,925.3766667	3,982,520.96933333	1,567,346.429	536,265.1585
A0A067P5W8	GT24	Glycosyltransferase family 24 protein	Y	48,176.3044	544,094.1723	253,305.0977	348,904.692666667
A0A067P7X8	GH105	Glycoside hydrolase family 105 protein	Y	29,812,627.2533333	25,171,134.76	12,159,050.3666667	21,464,407.3833333
A0A067P8J0	GT48	1,3-beta-glucan synthase	N	279,285.116566667	1,133,638.01133333	590,600.269866667	151,762.7827
A0A067P9S7	GT4	Glycosyltransferase family 4 protein	N	33,378,855.0866667	13,535,715.7066667	27,846,050.02	10,315,850.4533333
A0A067PAK9	GH43	Glycoside hydrolase family 43 protein	N	14,733.40841	543,993.021933333	85,749.15857	14,733.40841
A0A067PCV4	GH13	Alpha-amylase	Y	217,508.8117	148,575.9674	178,161.0273	157,544.848753333
A0A067PE74	GH31	Glycoside hydrolase family 31 protein	Y	260,600.315933333	23,874.92206	90,987.84162	23,874.92206
A0A0F6R9C3	AA1	Laccase A	Y	4,092,911.83733333	11,578,570.09	12,466,733.3766667	6,823,822.44233333
A0A2S1Q3T7	GT20	Alpha, alpha-trehalose-phosphate synthase (UDP-forming)	N	7,161,878.82433333	1,439,578.51166667	2,626,963.73133333	643,211.974533333
A0A482GMN1	AA2	Peroxidase	Y	30,817,611.0333333	13,001,488.19	28,196,940.2466667	23,719,528.7833333
A0A482GQ69	GH37	Trehalase	N	1,444,615.75266667	218,731.31909	51,030.5576433333	465,147.892066667

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A total of 66 proteins were annotated, with no CBM identified among the six categories of carbohydrate-active enzymes. The most abundantly expressed protein was GH7 glucanase (A0A067NSK0), which demonstrated the highest expression level in sugarcane bagasse. Its expression in corn cobs and cotton seed shells showed comparable levels, while significantly lower expression was observed in glucose medium compared to the other three substrates. Notably, this glucanase (A0A067NSK0) functions as a cellulase. Furthermore, the GH10 beta-xylanase (A0A067P3W7) exhibited relatively high expression levels compared to other proteins, with substantially enhanced expression in sugarcane bagasse compared to the other three media, while maintaining similar expression levels in corn cobs and cotton seed shells substrates. The GH72 family 1,3-beta-glucanosyltransferase (A0A067NTS1) showed elevated expression, particularly in cotton seed shells, where its expression markedly exceeded that in other substrates. Similarly, the GH3 glycoside hydrolase family 3 protein (A0A067N9V0) displayed higher expression in cotton seed shells. The CE15 carbohydrate esterase family 15 protein (A0A067NIA1) was predominantly expressed in sugarcane bagasse substrates.

The above results demonstrate that cellulases are dominant among the secreted carbohydrate-active enzymes. F. filiformis expresses a wide range of proteins involved in lignocellulosic substrate degradation. Proteomics analysis was conducted to identify predominant lignocellulose generated during this process and investigate substrate-specific enzymatic responses. Analysis of carbohydrate hydrolase expression profiles reveals that F. filiformis secreted functionally distinct CAZymes across different substrates, which serves as the underlying mechanism for the differential expression patterns of these proteins.

3.2. GO Functional Classification of Differential Proteins

Gene Ontology (GO) is a standardized framework for protein functional categorization. To elucidate the differences in secretory proteins between the three substrates and the glucose control group, differential proteins were subjected to GO functional annotation. The results are presented in Figure 2. Analysis reveals that the 1104 identified differential proteins were predominantly associated with biological processes, including cellular processes (685 proteins) and metabolic processes (664 proteins). For molecular functions, the majority were classified under catalytic activity (614 proteins) and binding (556 proteins). Regarding cellular components, the proteins were primarily localized to cellular anatomical entities (550 proteins) and protein-containing complexes (230 proteins).

Comparative GO functional clustering analysis between glucose and corn cobs substrates reveals enrichment of 625 differentially expressed proteins across 24 secondary terms, accounting for 56.61% of the total differential proteins. For glucose versus sugarcane bagasse, 596 differentially expressed proteins were enriched in 27 secondary terms, representing 53.99% of the total differential proteins. In glucose versus cotton seed shells substrates, 703 differentially expressed proteins were clustered into 23 secondary terms, constituting 63.68% of the total differential proteins. Shared enrichment analysis identifies 21 common GO terms across all comparisons. The most prominent term was cellular process, with 685 differential proteins (62.05% of total), followed by metabolic process (664 proteins, 60.14%), catalytic activity (614 proteins, 55.62%), binding (566 proteins, 51.27%), and cellular anatomical entity (550 proteins, 49.82%).

Figure 3 displays the top 23 significantly enriched GO terms for upregulated differential proteins and the top 23 enriched GO terms for downregulated differential proteins. All 46 GO terms met the significance threshold (p-adjust < 0.1). Comparative analysis reveals similar distribution patterns of differential proteins across the three substrates (sugarcane bagasse, cotton seed shells, and corn cobs) compared to the glucose control. Notably, sugarcane bagasse and cotton seed shells groups exhibited parallel modulation patterns. Within the catalytic activity category, both substrates showed higher enrichment of downregulated proteins than upregulated counterparts, suggesting potential functional suppression of these enzymes. Conversely, the corn cobs substrate demonstrated an inverse trend with predominant upregulation of catalytically active proteins, indicating functional potentiation. Figure 4 presents the GO enrichment analysis results for DEPs from the three substrates compared to the glucose substrate. As shown in Figure 4A, the results indicate that in the SCB vs. Glu group, the most significantly enriched terms were “non−membrane−bounded organelle” and “intracellular non−membrane−bounded organelle”. As shown in Figure 4B, in the CSS vs. Glu group, “ribosome biogenesis” was the most enriched term, while “protein dimerization activity” was most enriched in the CC vs. Glu group (Figure 4C). Collectively, these results indicate that the DEPs induced by different substrates differ in their functional distribution and enriched pathways, revealing F. filiformis’ adaptability to different substrates for growth.

GO analysis collectively demonstrates that differentially expressed proteins during substrate degradation are regulated by diverse cellular processes, particularly metabolic pathways. These findings highlight the multifaceted regulatory mechanisms governing lignocellulosic decomposition.

CSS: cotton seed shells, SCB: sugarcane bagasse, CC: corn cobs.

3.3. KEGG Pathway Enrichment Analysis of Differentially Expressed Proteins

Differentially expressed proteins with counts >1 were ranked by −log₁₀ p-value, and the top 20 enriched KEGG pathways were visualized as a bubble plot (Figure 5). Figure 5 shows the significantly enriched metabolic pathways. A total of 380 DEPs were enriched between glucose and corn cob substrates and annotated to 91 pathways. The KEGG enrichment analysis bubble chart reveals that the pathways with higher enrichment scores include arginine and proline metabolism (three upregulated, eight downregulated), cysteine and methionine metabolism (three upregulated, eight downregulated), and phenylalanine metabolism (five upregulated, two downregulated). Between glucose and cotton seed shells substrates, 405 DEPs were enriched and annotated to 91 pathways. The KEGG enrichment analysis bubble chart indicates that the pathways with higher enrichment scores include phenylalanine metabolism (two upregulated, five downregulated) and tryptophan metabolism (five upregulated, five downregulated). A total of 357 DEPs were annotated to 82 pathways between glucose and sugarcane bagasse substrates. The KEGG enrichment analysis bubble chart shows that the pathways with higher enrichment scores include galactose metabolism (four upregulated, four downregulated), ribosome (two upregulated, forty-one downregulated), and starch and sucrose metabolism (nine upregulated, nine downregulated).

When compared to glucose substrates, the annotation results of DEPs in corn cobs and cotton seed shells were predominantly enriched in amino acid metabolic pathways, whereas sugarcane bagasse substrates showed DEPs primarily associated with carbohydrate metabolism. Metabolomic analysis of F. filiformis cultivated on substrates with varying C/N ratios reveals that differentially abundant metabolites were significantly enriched in amino acid biosynthesis and metabolism pathways, with corresponding variations in fruiting body amino acid content [28]. Targeted metabolomic profiling of Auricularia heimuer fruiting bodies grown on sawdust, cotton seed shells, and corn cobs demonstrates that differential metabolites were enriched in amino acid synthesis and metabolism pathways. Notably, the content of nine essential amino acids in A. heimuer cultivated on cotton seed shells was significantly higher than those grown on sawdust and corn cobs, indicating that cotton seed shells and corn cob matrices effectively enhance amino acid accumulation [29]. These findings suggest that corn cobs and cotton seed shells may similarly promote amino acid biosynthesis in F. filiformis.

We observed that within the phenylalanine metabolism pathway, corn cobs and cotton seed shells exhibited enrichment of identical protein categories among DEPs. However, amine oxidase (A0A067NX95), annotated as a copper amine oxidase (CuAO), was downregulated in corn cobs and upregulated in cotton seed shells, with no significant enrichment detected in sugarcane bagasse. Copper amine oxidases (CuAOs) are emerging as physiologically critical enzymes due to their roles in plant growth, differentiation, and defense responses to biotic and abiotic stresses [30]. In plants, CuAO stress adaptation primarily relies on the catalytic products of polyamine catabolism—γ-aminobutyric acid (GABA) and H₂O₂ [31].

Aspartate aminotransferase (A0A067P1S4) exhibited downregulated expression in corn cobs while being upregulated in cotton seed shells and sugarcane bagasse. Aspartate aminotransferase (AAT), also known as glutamic–oxaloacetic transaminase (GOT), catalyzes a reversible transamination reaction between glutamate (Glu) and oxaloacetate (OAA) to generate aspartate (Asp) and 2-oxoglutarate (OG) in a pyridoxal 5′-phosphate (PLP)-dependent manner. This enzyme is ubiquitously distributed across animals, plants, and microorganisms and plays a pivotal role in nitrogen (N) shuttling between Glu and Asp in all organisms [32]. AAT is critical for fungal nitrogen assimilation and carbon–nitrogen balance and is implicated in plant abiotic stress response pathways [33].

Phenylalanine ammonia-lyase (PAL, A0A4P8P9Z1) exhibited upregulated expression in corn cobs while showing downregulated expression in cotton seed shells and sugarcane bagasse. As the initial enzyme catalyzing the biosynthesis of precursors for various secondary metabolites (including flavonoids), PAL plays crucial roles in regulating plant growth, development, and responses to biotic and abiotic stresses [34,35]. Omics analysis of F. filiformis fruiting body development reveals a significant positive correlation between PAL gene expression levels and flavonoid content. This suggests PAL’s involvement in flavonoid biosynthesis and metabolism and its regulatory function in the growth and developmental processes of F. filiformis [36].

This study observed significant differences in the expression patterns of key enzymes within the phenylalanine metabolism pathway across the three lignocellulosic substrates (corn cobs, cotton seed shells, and sugarcane bagasse). Although the specific types of differentially abundant proteins enriched in this pathway were consistent across substrates, the direction of change in expression abundance for A0A067NX95 (K00276, amine oxidase, and CuAO) and A0A067P1S4 (K14454, aspartate aminotransferase, and AAT) was substrate-specific. The distinct expression profiles of these key enzymes suggest that phenylalanine metabolism may play an important role in F. filiformis’ adaptation to utilizing different lignocellulosic substrates. This demonstrates the substantial influence of substrate composition on the fungal metabolic network. Furthermore, these findings provide insights into how F. filiformis might regulate primary and secondary metabolism to adapt to diverse environments during the valorization of agricultural waste. Comparative proteomic analysis between sugarcane bagasse and glucose substrates reveals significant enrichment of proteins associated with the ribosome, starch and sucrose metabolism and galactose metabolism. Within the starch and sucrose metabolism pathway, several enzymes exhibited upregulated expression, including glucanase (A0A067NSK0), phosphoglucomutase (A0A067NHY6), and glucose-6-phosphate isomerase (A0A067NV87). Notably, phosphoglucomutase (PGM), UDP-glucose pyrophosphorylase (UGPP), and phosphomannose isomerase (PMI) have been identified as critical enzymes in fungal polysaccharide biosynthesis, with glucose-6-phosphate isomerase (GPI), playing pivotal roles in both glycolysis and gluconeogenesis [37]. Omics-based studies on Ganoderma lucidum polysaccharide production further identified key glycosyltransferases and glycoside hydrolases involved in polysaccharide synthesis. Intriguingly, polysaccharide yields in G. lucidum varied significantly when cultivated on glucose versus xylose as carbon sources [38]. Therefore, the observed upregulation of these carbohydrate metabolism enzymes suggests a potential enhancement of carbon flux towards pathways that could support polysaccharide synthesis in F. filiformis grown on sugarcane bagasse.

The upregulated expression of 1,3-beta-glucan synthase (A0A067P8J0) is associated with the synthesis of β-1,3-glucan, a key polysaccharide component unique to fungal cell walls. This polysaccharide is synthesized by the β-1,3-glucan synthase complex and plays a critical role in polysaccharide biosynthesis. Previous studies have demonstrated that silencing or overexpression of the gene encoding 1,3-β-glucan synthase in Ganoderma lucidum and Cordyceps militaris affects polysaccharide synthesis [39]. Similarly, the upregulated expression of this protein in sugarcane bagasse substrates may also influence polysaccharide synthesis and, consequently, alter the composition of cell wall components.

Trehalose not only serves as a storage carbohydrate but also functions as a protective factor. Trehalose-6-phosphate synthase (Tps1) catalyzes the formation of trehalose-6-phosphate (T6P) from uridine diphosphate glucose (UDPG) and α-glucose-6-phosphate (α-Glc-6-P). Subsequently, T6P is dephosphorylated to trehalose through the action of trehalose-6-phosphate phosphatase (Tps2). The catabolism of trehalose is catalyzed by trehalase [40]. In this study, alpha, alpha-trehalose-phosphate synthase (A0A2S1Q3T7), and trehalase (A0A482GQ69) were found to be downregulated. Referring to the trehalose metabolic pathway summarized in the study [21], the DEP enrichment results in this research indicated that PGM was upregulated. This suggests that trehalose may be converted to glucose-6-phosphate, implying that the accumulation of trehalose might be affected in sugarcane bagasse. This altered protein profile may impact cellular trehalose pools, which are critical for stress tolerance in F. filiformis.

In the galactose metabolism pathway, α-galactosidase (A0A067NW26) exhibited upregulated expression. α-galactosidase is a member of the hemicellulase enzyme system. Synergistic effects have been reported among members of the hemicellulase system, such as between galactosidase and mannanase [41]. Zhu et al. [42] demonstrated that when the crude enzyme extract of the consortium EMSD5 (primarily hemicellulose-degrading enzymes) was combined with commercial cellulase preparations in a 3:1 ratio to treat corn stover, the maximum synergistic coefficient reached 1.96, with cellulose and xylan conversion rates of 38.3% and 18.7%, respectively. The degradation of lignocellulose requires synergistic action among various enzymes. The cross-linked network of lignocellulose limits enzyme function, but this limitation can be partially overcome through synergistic effects [43]. In this study, the synergistic effect of the lignin-degrading enzyme system played a certain role in the adaptation of F. filiformis to the sugarcane bagasse.

Collectively, proteomic analysis reveals that the central metabolic pathways of F. filiformis exhibit distinct substrate-dependent characteristics when grown on three different substrates: corn cobs, cotton seed shells, and sugarcane bagasse. Specifically, these differences are mainly reflected in processes such as amino acid metabolism and carbohydrate metabolism.

In-depth analysis of proteins enriched in metabolism pathways reveals significant clustering patterns among DEPs, as illustrated in Figure 6. Comparative DEPs between sugarcane bagasse and glucose substrates (Figure 6A) identified two prominently enriched proteins: glucanase (A0A067NSK0) and ATP synthase subunit beta (A0A067NZS0). The ATP synthase subunit beta, a critical component of the ATP synthase complex, participates in oxidative phosphorylation by catalyzing ATP synthesis and hydrolysis. This enzyme comprises the F1 and F0 subunits, with the β-subunit harboring catalytic sites essential for ATP production [44]. The abundance of ATP synthase subunit beta was significantly upregulated in sugarcane bagasse, indicating that, compared to glucose, the components of the mitochondrial ATP synthesis mechanism were more abundant under sugarcane bagasse. Glucanase (A0A067NSK0), a GH7 glycoside hydrolase, functions as an endo-β-1,4-glucanase that randomly hydrolyzes β-1,4-glycosidic bonds in amorphous cellulose regions [45]. Notably, prior studies on Aspergillus fumigatus demonstrated that heterologous expression of the GH7-family endoglucanase Af-EGL7 (encoded by Afu6g01800) in E. coli resulted in a 2500-fold induction when cultured on sugarcane exploded bagasse (SEB), highlighting its exceptional cellulolytic efficiency [46]. The abundance of A0A067NSK0 (glucanase, GH7) was significantly upregulated on both sugarcane bagasse and cotton seed shells. (It was also one of the differentially expressed proteins on the corn cobs; see Figure 6C.) This suggests that on lignocellulosic substrates with cellulose as the main component, F. filiformis may have enhanced the expression of genes encoding specific endoglucanases, which is consistent with its need to degrade lignocellulosic substrates. Comparative analysis of DEPs between corn cobs and glucose substrates reveals prominent enrichment of two proteins: ATP synthase subunit beta (A0A067NZS0) and phosphopyruvate hydratase (A0A067P4U9), as shown in Figure 6C. Phosphopyruvate hydratase catalyzes the dehydration of 2-phosphoglycerate (2-PG) to phosphoenolpyruvate (PEP), representing the penultimate step in the glycolytic pathway [47].

Comparative DEP analysis between cotton seed shells and glucose substrates reveals significant enrichment of two proteins in Figure 6B: glucanase (A0A067NSK0) and glyceraldehyde-3-phosphate dehydrogenase, GAPDH (D0VBH9). GAPDH, a pivotal glycolytic enzyme, catalyzes the oxidative phosphorylation of glyceraldehyde-3-phosphate (G-3-P) to produce 1,3-bisphosphoglycerate. When F. filiformis was grown on the three lignocellulosic substrates, the abundance of some key proteins involved in the glycolysis and oxidative phosphorylation pathways generally showed an increasing trend compared to those grown on glucose substrates.

In all three lignocellulosic substrates, the expression levels of nucleoside diphosphate kinase, NDPK (A0A067PBA8), were consistently lower compared to those observed in glucose-based media. NDPK plays a critical role in maintaining nucleotide homeostasis by transferring the γ-phosphate group from nucleoside triphosphates (NTPs) to nucleoside diphosphates (NDPs), thereby regenerating NTPs—a process essential for cellular energy metabolism and nucleic acid biosynthesis. These substrate-dependent variations in enzyme expression further modulate energy metabolism dynamics. Similarly, A0A067NI01 (UTP-glucose-1-phosphate uridylyltransferase), also known as UDP-glucose pyrophosphorylase (UGPase), exhibited reduced expression across all substrates relative to glucose. UGPase catalyzes the conversion of UTP and glucose-1-phosphate (G-1-P) to UDP-glucose (UDP-Glc) and pyrophosphate (PPi), with UDP-Glc serving as a pivotal precursor for glycogen and starch biosynthesis. In the comparison of differentially expressed proteins, the consistent downregulation of NDPK and UGPase suggests that the abundance of proteins related to nucleotide metabolism and sugar nucleotide synthesis pathways in F. filiformis grown on lignocellulosic substrates differs from that in those grown on glucose substrates.

Clustering analysis (Figure 6) and the differential protein abundance data clearly demonstrate that when F. filiformis is cultivated on three distinct lignocellulosic substrates (sugarcane bagasse, corn cobs, and cotton seed shells), its proteome undergoes significant changes. This is characterized by increased abundance of proteins associated with energy metabolism, decreased abundance of proteins involved in nucleotide metabolism and sugar nucleotide synthesis, and increased abundance of cellulose-degrading enzymes. These observed differences in protein abundance provide important molecular insights into the metabolic strategies employed by F. filiformis to adapt to different lignocellulosic matrices. However, due to the inherent limitations of proteomics, further investigation into metabolic products is essential to determine how these specific protein abundance changes translate to actual substrate degradation efficiency.

Constrained by the experimental design, the proteomic results of F. filiformis across different substrates only provide preliminary indications of the molecular basis underlying its environmental adaptation. The identified GH7 glucanase (A0A067NSK0) may serve as a candidate for future development of lignocellulose-degrading enzymes. The proteomic data tentatively suggest that substrate type could influence yield and quality, but independent replication and functional validation are required. These exploratory findings provide preliminary insights into substrate-adaptation strategies, thereby laying the groundwork for future research on agricultural waste valorization and high-value metabolite production.

4. Discussion

It is well established that the lignocellulolytic enzyme profiles secreted by fungi are strongly influenced by the type and composition of substrates [48]. Thus, culturing fungi on multiple substrates enables a broader understanding of their lignocellulose degradation capabilities. In this study, F. filiformis was cultivated on four distinct substrates to elucidate the composition of its core enzymatic systems and degradation strategies for lignocellulose breakdown.

In this study, comparative analysis of the secretory protein of F. filiformis across four substrates via the dbCAN database identified 66 proteins. Among the CAZymes, glycoside hydrolases (GHs; EC 3.2.1.) represented the most abundant group, with 41 GH proteins identified out of the 66 annotated proteins. Notably, a GH7 glucanase (A0A067NSK0) exhibited the highest expression level and played a predominant role in cellulose degradation. Comparative genomic analysis of Flammulina velutipes var. lupinicola reveals that its genome encodes a suite of cellulolytic enzymes (GH5, -6, -7, -9, and -12) [49], yet the protein from GH7 was relatively less annotated. This contrast underscores the high application potential of the GH7 glycoside hydrolase identified in this study, which demonstrates exceptional functional specificity and catalytic efficiency in lignocellulose deconstruction.

SCHMITZ et al. [50] reported that the synergistic degradation of three substrates (corn, wheat bran, and oats) by multiple ligninolytic and cellulolytic enzymes involves substrate-specific modes of action targeting both the structural backbone and recalcitrant components, such as lignin, with varying conversion efficiencies and cooperative interactions. These findings align with observations in the present study, wherein F. filiformis degrades different substrates through a complex enzyme system. In sugarcane bagasse, F. filiformis primarily utilizes glucanase (A0A067NSK0) and hemicellulase beta-xylanase (A0A067P3W7). Conversely, cotton seed shells predominantly involve two cellulases: glucanase (A0A067NSK0) and a GH enzyme (A0A067N9V0). Notably, corn cobs degradation relies chiefly on glucanase (A0A067NSK0). To access bioavailable carbon from complex lignocellulosic matrices, fungi enhance their capacity to utilize recalcitrant carbon sources by secreting copious extracellular enzymes [51], which likely explains the differential cellulase expression levels observed across the three substrates in this study.

In this study, the number of ligninolytic enzymes identified through functional enrichment was limited, with the primary annotated lignin-degrading enzymes being AA1 laccase A (A0A0F6R9C3) and AA2 peroxidase (A0A482GMN1). A genome-wide and global gene expression analysis of the model mushroom Flammulina velutipes reveals differential expression levels of AA genes across developmental stages and tissues. Notably, six AA genes (one AA1, one AA2, two AA3, and two AA5) exhibited consistently higher overall expression levels than other AA genes in all stages and tissues [52], aligning with the findings of this work. Previous studies on lignin-degrading model fungi have demonstrated that nutrient restriction of carbon source and nitrogen stimulates the synthesis of ligninolytic enzymes [24], which explains why laccase and peroxidase were most enriched on corn cobs. Enzymes involved in the degradation of pectin, galactomannan, and xyloglucan were detected at low diversity and abundance, consistent with the paucity of genes encoding these enzymes in the fungal genome [49].

Proteomic analysis of the secretome reveals significant enrichment of DEPs in energy metabolism and carbohydrate metabolism pathways. In sugarcane bagasse substrates, substantial enrichment was observed not only for the ATP synthase subunit beta (A0A067NZS0) but also for other metabolism-related DEPs, whereas no significant enrichment of these proteins occurred in the other two substrates. This suggests distinct regulatory mechanisms in ATP metabolism during F. filiformis cultivation on sugarcane bagasse. In contrast, cotton seed shells and corn cobs exhibited increased enrichment of key glycolytic enzymes, including GAPDH and phosphopyruvate hydratase, indicating altered glycolysis flux. The cell walls of filamentous fungi are primarily composed of chitin and β-1,3-glucans, with CAZyme-associated regulatory genes playing critical roles in cell wall biosynthesis [53]. Previous studies have identified phosphoglucomutase (PGM), UDP–glucose pyrophosphorylase (UGPase), and phosphomannose isomerase (PMI) as key enzymes in fungal polysaccharide synthesis pathways [54]. Notably, reduced enrichment of UGPase was observed across substrates. The abundance of UGPase is reduced on the substrate, and its downregulation may affect the process of cell wall remodeling.

Proteomic analysis of F. filiformis cultivated on diverse substrates preliminarily reveals molecular foundations underlying its environmental adaptability, holding significant biotechnological implications. The identified GH7 glucanase (A0A067NSK0) exhibits potential for developing novel lignocellulose-degrading enzymes. These initial findings provide exploratory clues for optimizing culture medium formulations to enhance yield and quality, serving as a reference for further elucidating substrate-adaptation strategies, and may inform future efforts to valorize agricultural waste and produce high-value metabolites.

This study provides a preliminary profiling of the secretome of F. filiformis cultivated on four different substrates. Two limitations should be noted. First, although the fungal mycelia were derived from independent culture media, all replicates were processed within a single experimental batch. This design constitutes technical replication rather than independent biological replication across multiple batches or temporal stages. Consequently, the identified substrate-specific proteomic patterns should be interpreted as preliminary and hypothesis-generating, requiring validation through true biological replication. Future studies should verify these findings across multiple cultivation batches and timepoints to establish robust biological patterns. Additionally, while this work focuses on proteomic responses, data on the growth performance of the strain across different substrates were not incorporated. Integrating such physiological parameters with multiomics approaches in subsequent research will further validate and extend the applicability of our conclusions.

5. Conclusions

Quantitative proteomic analysis of the secretory proteome from F. filiformis cultivated on four substrates identifies the GH7 family glucanase (A0A067NSK0) as an important enzyme in cellulose degradation. On the sugarcane bagasse, F. filiformis secretes higher levels of cellulases and hemicellulases, which act synergistically to degrade lignocellulose. On the cotton seed shells, multiple cellulases work together to degrade cellulose. On the corn cobs, the GH7 family glucanase is the predominant cellulase. This study provides exploratory evidence for the resource utilization of lignocellulolytic enzymes and offers preliminary guidance for optimizing F. filiformis cultivation substrates. However, several limitations should be acknowledged: the observed differential expression patterns remain suggestive rather than conclusive regarding functional contributions. Future work should quantify the activity of key enzymes to verify their contribution to lignocellulose degradation, elucidate the synergistic mechanisms of the enzyme system, and systematically investigate the structure–activity relationship between substrate composition and the secretome profile to refine our understanding of the fungus’ lignocellulose-degrading machinery.