Changes in Microbial Diversity During Dictyophora indusiata Mycelium Regression Period

Abstract

Dictyophora indusiata cultivation is severely impeded by premature hyphal regression. This study elucidates the spatiotemporal dynamics of mycelial regression and associated microbial succession in both substrate and soil matrices across progressive regression stages (CK: normal growth; S1: initial recession; S2: advanced recession; S3: complete recession). Microscopic analysis revealed preferential mycelial regression in the substrate, preceding soil regression by 1–2 stages. High-throughput sequencing demonstrated significant fungal community restructuring, characterized by a sharp decline in Phallus abundance (substrate: 99.7% → 7.0%; soil: 78.3% → 5.5%) and concomitant explosive proliferation of Trichoderma (substrate: 0% → 45.2%; soil: 0.1% → 55.3%). Soil fungal communities exhibited a higher richness (Chao1, p < 0.05) and stability, attributed to functional redundancy (e.g., Aspergillus, Conocybe) and physical protection by organic–mineral complexes. Conversely, substrate bacterial diversity dominated, driven by organic matter availability (e.g., the Burkholderia–Caballeronia–Paraburkholderia complex surged to 59%) and optimized porosity. Niche analysis confirmed intensified competition in post-regression soil (niche differentiation) versus substrate niche contraction under Trichoderma dominance. Critically, Trichoderma overgrowing was mechanistically linked to (1) nutrient competition via activated hydrolases (e.g., Chit42) and (2) pathogenic activity (e.g., T. koningii causing rot). We propose ecological control strategies: application of antagonistic Bacillus subtilis (reducing Trichoderma by 63%), substrate C/N ratio modulation via soybean meal amendment, and Sphingomonas–biochar soil remediation. This work provides the first integrated microbial niche model for D. indusiata regression, establishing a foundation for sustainable cultivation.

1. Introduction

Dictyophora indusiata Fisch. is a nutrient-rich, large edible fungus prized for its crisp texture, elegant morphology, distinctive aroma, and high nutritional value [1]. Its polysaccharides are particularly valued for enhancing immune function [2], regulating gut microbiota [3], and alleviating obesity [4]. Key species include D. indusiata (long-skirt bamboo fungus), Dictyophora rubrovolvata (red-veiled bamboo fungus), Phallus echinovolvatus (spiny-volva stinkhorn), and Fructificatio Dictyophorae Multi-coloris (yellow-skirt bamboo fungus). Among these, D. indusiata is a renowned edible and medicinal species within the genus [5]. D. indusiata is primarily concentrated in regions south of the Yangtze River, including Fujian, Sichuan, and Guizhou. Fujian, Jiangxi, and Sichuan represent the dominant production areas, with a combined output of 188.1 thousand tons in 2021, accounting for 97.12% of China’s total D. indusiata yield. The anomalous “regression” phenomenon in D. indusiata mycelium refers to the disappearance and degeneration of mycelia within the substrate and soil during the mycelial growth phase. This phenomenon subsequently prevents primordium formation or disrupts fruiting body development after primordium initiation, ultimately leading to yield reduction or total crop failure.

Early studies by Wu Shaofeng et al. [6] suggested ammonia toxicity as the primary cause of mycelial atrophy. Subsequent research on soil-cultivated fungi revealed that soil microbiota significantly impact their growth. Microbial influences on soil-cultivated edible fungi may lead to a reduced yield and quality; these effects primarily manifest in competition, infection, and antagonism. Fungi of the genus Trichoderma spp. are the primary infectious agents of green mold disease in edible fungi, representing highly pathogenic competitive contaminants [7]; Mycogone perniciosa causes brown blotch disease in Agaricus bisporus [8]; Fusarium incarnatum induces stipe rot in Morchella spp. [9].

Microorganisms play critical roles in forming selective substrates, providing nutrients, stimulating growth and fruiting body formation, and defending against pathogens. For example, adding a tannin–saponin-degrading microbial agent containing Bacillus amyloliquefaciens and Aspergillus awamori to Camellia oleifera shells improves cultivation substrates and promotes rapid fungal growth [10]; Sun et al. [11] isolated the plant growth-promoting rhizobacterium Serratia odorifera HZSO-1 from mushroom cultivation material, which enhances the growth of Hypsizygus marmoreus, shortens the primordiation period by 3–4 d, and increases fruiting body yield by 12%. Bacillus subtilis [12], Bacillus velezensis [13], Bacillus amyloliquefaciens [14], and Aureobasidium pullulans [15] exhibit antagonistic effects against Trichoderma, effectively controlling green mold disease. Pseudomonas putida can attach to A. bisporus mycelia and produce volatile organic compounds (VOCs) [16], thereby stimulating mycelial growth and primordium initiation. Notably, many researchers consider a reduced mass concentration of VOCs as one environmental signal inducing fruiting in A. bisporus [17].

This study sampled soil and substrate from regression-affected areas in Yibin across different regression stages. High-throughput sequencing of microbial diversity, combined with microscopic analysis, elucidated regression patterns and causes, informing scientific control strategies for this phenomenon.

2. Materials and Methods

2.1. Experimental Materials

Soil samples (containing D. indusiata mycelium) and substrate (bamboo sawdust) were derived from the Daguan Experimental Base of the Yibin Academy of Agricultural Sciences. The cultivated bamboo fungus species was D. indusiata (long-skirt bamboo fungus).

Samples were collected using the five-point sampling method (covering central and four corner positions of the cultivation area). Sterilized tools included alcohol flame-sterilized stainless steel core samplers (100 cm³) and soil augers (3 replicates per point). Sample processing; homogenization: Composite samples were mixed via the quartering method using sterile gloves. Packaging: Distributed into sterile 1.5 mL centrifuge tubes. Total per sample: Twelve tubes (representing 20 g composite sample). Preservation: Flash-frozen in liquid nitrogen. Transferred to −80 °C ultra-low-temperature freezer (to prevent DNA degradation). Sampling time, number, and mycelium state (mycelium in soil and substrate) were divided according to the standard of mycelium regression periods of edible fungi [18] as shown in the following table (

Table 1. The number of soil and substrate samples, the sampling time, and the mycelium state referred to by the sampling standard.

Substrate Sample ID	Soil Sample ID	Sampling Time	Mycelial Status
CK	CKTU	2024/12	Normal growth stage
S1	S1TU	2025/1	Initial recession stage (>70% mycelium retained)
S2	S2TU	2025/2	Mid-recession stage (30–50% mycelium retained)
S3	S3TU	2025/3	Complete recession stage (<5% mycelium retained)

).

2.2. Experimental Methods

2.2.1. Mycelial Dynamic Microscopy

Instrument: Leica DVM6 3D (Leica Microsystems Ernst-Leitz-Straße17-37 35578 Wetzlar, Germany) digital microscopy system. Sample pretreatment: Fresh samples. Observation: Five random fields of view per sample were examined at 500× magnification to record mycelial morphology and coverage.

2.2.2. DNA Extraction and Quality Control

Kit: MagBeads FastDNA Kit for Soil (116564384; MP Biomedicals, CA, USA). Critical step: SLX-Mlus lysis buffer was added to eliminate humic acid interference [19]. Mechanical lysis: Liquid nitrogen grinding followed by vortex oscillation (3 × 30 s). QC standards: Nanodrop: A₂₆₀/A₂₈₀ = 1.8–2.0, concentration > 20 ng/μL. Gel electrophoresis: 1.2% agarose gel, dominant band > 5000 bp. Storage: −20 °C (short-term)/−80 °C (long-term). Samples were repeated three times.

2.2.3. PCR Amplification and Sequencing

The target region, primer sequence, amplification system, and procedure are shown in Table 2. Sequencing: Perseno Biotechnology Company was entrusted to complete the sequencing (Shanghai, China).

DNA denoising was carried out by using the Data2 (https://github.com/benjjneb/dada2, accessed on 1 August 2025) [20] software platform: firstly, the primer fragment of the qiime cutadapt trim-paired excision sequence is called, and the sequence of unmatched primers is discarded; Then DADA2 is called by qiime dada2 denoise-paired for quality control, denoising, and mosaic and chimera removal. The above steps are analyzed separately for each library. After denoising all the libraries, ASV feature sequences and ASV tables are merged, and singletons ASVs are removed.

Table 2. PCR amplification region, primer sequence, system, and program setting.

	Target Region	Primer Sequence	Amplification System	Procedure
Fungal	ITS1 region is about 300 bp in length	ITS5: 5′-GGAAGTAAAAGTCGTAACAAGG-3′ ITS2: 5′-GCTGCGTTCTTCATCGATGC-3′ [21]	DNA template: 2 μL 5× GC buffer: 5 μL dNTPs (2.5 mM): 2 μL Forward/Reverse primers (10 μM): 1 μL each Q5 High-Fidelity DNA Polymerase: 0.25 μL ddH2O 8.75 μL	98 °C for 2 min, predenaturation; 30 cycles of [98 °C for 15 s, 55 °C for 30 s, 72 °C for 30 s]; 72 °C for 5 min
Bacterial	V3V4 hypervariable region, with a length of about 460 bp.	338F: 5′-ACTCCTACGGGAGGCAGCAG-3′ 806R: 5′-GGACTACHVGGGTWTCTAAT-3′ [22]	DNA template: 2 μL 2× KAPA HiFi HotStart Mix: 12.5 μL Forward/Reverse primers (5 μM): 1 μL, ddH2O 8.5 μL	98 °C for 3 min, predenaturation; 25 cycles of [95 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s]; 72 °C for 5 min

Table 2. PCR amplification region, primer sequence, system, and program setting.

	Target Region	Primer Sequence	Amplification System	Procedure
Fungal	ITS1 region is about 300 bp in length	ITS5: 5′-GGAAGTAAAAGTCGTAACAAGG-3′ ITS2: 5′-GCTGCGTTCTTCATCGATGC-3′ [21]	DNA template: 2 μL 5× GC buffer: 5 μL dNTPs (2.5 mM): 2 μL Forward/Reverse primers (10 μM): 1 μL each Q5 High-Fidelity DNA Polymerase: 0.25 μL ddH2O 8.75 μL	98 °C for 2 min, predenaturation; 30 cycles of [98 °C for 15 s, 55 °C for 30 s, 72 °C for 30 s]; 72 °C for 5 min
Bacterial	V3V4 hypervariable region, with a length of about 460 bp.	338F: 5′-ACTCCTACGGGAGGCAGCAG-3′ 806R: 5′-GGACTACHVGGGTWTCTAAT-3′ [22]	DNA template: 2 μL 2× KAPA HiFi HotStart Mix: 12.5 μL Forward/Reverse primers (5 μM): 1 μL, ddH2O 8.5 μL	98 °C for 3 min, predenaturation; 25 cycles of [95 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s]; 72 °C for 5 min

2.2.4. Data Processing

Reference Database: The UNITE database (Release 8.3; https://unite.ut.ee/ accessed on 5 April 2025) [23] was used as the default reference for taxonomic assignment. Alpha Diversity Analysis: Preliminary data organization was performed in Microsoft Excel 2003 (standard edition). Alpha diversity indices were calculated based on rarefied feature tables. Taxonomic Composition Visualization: Bar plots depicting community composition at the genus level were generated using the circlize package (v0.4.16) in R (v4.3.3). These visualizations were based on feature tables after removal of singleton amplicon sequence variants (ASVs)/operational taxonomic units (OTUs). Beta Diversity Analysis: Beta diversity metrics were computed to assess microbial community dissimilarities between distinct habitats using QIIME 2 (2024.5 release). Principal Coordinate Analysis (PCoA) was performed in R (v4.3.3) with the ape (v5.8) and vegan (v2.6-6.1) packages, with results visualized as two-dimensional scatter plots. Venn Diagram Analysis: Venn diagrams were constructed in R (v4.3.3) to identify shared and unique ASVs/OTUs among sample groups. Each experimental group was treated as a distinct set, with ASVs/OTUs (present at >0 abundance across all samples) assigned to sets based on their presence/absence in group replicates.

3. Results

3.1. Microscopic Observation of Mycelial Decline

As illustrated in Figure 1A–D, the recession process of D. phalloidea mycelia exhibited distinct stages: Normal growth stage (Figure 1A): Substrate appeared yellow and was densely colonized by white mycelia. Abundant mycelia were also distributed throughout the soil. Initial recession stage (Figure 1B): A marked transition in substrate color from yellow to brown was observed. Sparse mycelial remnants persisted within the browned substrate, whereas no discernible changes occurred in soil mycelia. Advanced recession stage (Figure 1C): Substrate turned uniformly brown, with a complete absence of mycelia. Soil mycelia remained quantitatively stable with no visible reduction. Complete recession stage (Figure 1D): Both substrate and soil exhibited total mycelial depletion, with substrate maintaining a brown coloration. Key observation: Mycelial recession was initiated exclusively in the substrate; soil mycelia underwent recession only after complete substrate mycelial depletion, suggesting a sequential degradation pattern.

Microscopic observations of substrate–soil systems (Figure 2: CKTU/S1TU/S2TU/S3TU for soil; CK/S1/S2/S3 for substrate) revealed a significant spatiotemporal heterogeneity in mycelial recession, characterized by three distinct phases: Stage-Specific Dynamics S1 Phase (Initial Recession): No substantial structural changes were detected in mycelial networks of either substrate or soil. S2 Phase (Advanced Recession): Substrate: Preferential collapse of mycelial networks occurred, with complete disintegration of formerly integrated structures. Dense tan-colored aggregates fragmented into loose detritus, accompanied by numerous severed hyphal fragments. Soil: Exhibited a delayed response, where partial mycelia underwent yellowing (pigment shift from white to pale yellow), indicating incipient degradation. S3 Phase (Terminal Recession): Complete absence of mycelia was confirmed in both substrate and soil matrices. Key Scientific Findings, Spatial Priority: Mycelial degradation was initiated exclusively in the substrate (S2 phase), preceding soil mycelium recession. Morphological Biomarkers: Network disintegration and hyphal fragmentation in substrate serve as early indicators of systemic collapse. Delayed Soil Response: Soil mycelial degradation manifested primarily through chromatic transition (yellowing), occurring asynchronously with substrate collapse.

3.2. High-Throughput Sequencing Result Analysis

In this study, eight samples were taken from the substrate and soil of D. indusiata mycelium at different periods (each sample was repeated three times).

Table A3. Statistical table of sample sequencing quantity.

Sample ID	Input	Filtered	Denoised	Merged	Non-Chimeric	Non-Singleton
CKTU	109,776.33 ± 5629.93	102,697.00 ± 5182.25	94,873.33 ± 5763.75	67,921.66 ± 6893.47	62,097.33 ± 6290.94	61,371.66 ± 6333.50
CK	99,585.66 ± 27523.65	92,421.66 ± 26,020.03	91,326.66 ± 26,227.15	87,454.33 ± 27,545.90	78,003.00 ± 30,029.23	77,937.33 ± 30,075.59
S1TU	116,584.66 ± 1498.26	108,273.00 ± 1192.61	100,809.66 ± 2152.39	74,537.00 ± 4509.07	65,382.66 ± 4556.47	64,603.00 ± 4901.19
S1	103,829.33 ± 7107.30	95,390.33 ± 6547.03	94,180.33 ± 6244.19	90,153.00 ± 5241.89	68,152.66 ± 6389.29	68,045.33 ± 6427.77
S2 TU	105,704.66 ± 770.75	98,345.00 ± 1207.74	95,891.00 ± 1909.29	87,926.33 ± 8992.41	85,788.33 ± 10,503.46	85,629.00 ± 10,718.21
S2	87,332.66 ± 19,457.50	80,560.00 ± 17,796.59	79,445.33 ± 18,095.25	76,628.66 ± 19,182.23	58,473.66 ± 14,945.13	58,419.33 ± 14,693.78
S3TU	122,144.66 ± 4599.45	113,536.66 ± 4223.50	104,705.00 ± 3913.49	75,480.00 ± 3624.93	69,164.66 ± 3341.63	68,076.00 ± 3406.49
S3	106,029.00 ± 12,826.89	98,328.33 ± 11,937.87	94,693.62 ± 12613.84	88,382.66 ± 12,749.27	61,607.66 ± 11,362.59	61,427.66 ± 11,417.18

Note: The letters after the numbers of peers are significant differences (p ≤ 0.05; n = 3). Input is the sequence quantity that can match both forward and reverse primers in the original data; Filtered is the amount of data after removing low-quality sequences; Denoised is the amount of sequence data after denoising, that is, the effective sequence amount; Merged is the sequence quantity after splicing; Non-Chimeric is the sequence quantity after chimera removal, which is the high-quality sequence quantity; Non-Singleton is the sequence quantity after removing singletons.

shows that 2,603,510 high-quality sequences were extracted from the eight samples, including 1,289,535 sequences in soil and 1,313,975 sequences in substrate. In sparse curves of all samples (Figure A1A,B), when the flattening depth reaches about 2000 (fungi) and 40,000 (bacteria), all sample curves tend to be flat. The smoothness of the curve reflects the influence of sequencing depth on the diversity of observed samples, which shows that most fungi in the samples will be detected in this experiment.

3.3. Alpha Diversity Analysis

Five alpha diversity indices (Chao1, Simpson, Pielou’s evenness, Shannon, and Observed species) were employed to characterize microbial community shifts during Dictyophora’s mycelial recession (Figure 3A and

Table A1. Fungal alpha diversity analysis table.

Sample Name	Chao1	Simpson	Pieloue	Shannon	Observed Species
CK	452.29 ± 39.14 a	0.967 ± 0.002 a	0.73 ± 0.014 a	4.455 ± 0.147 a	450.000 ± 37.723 a
CKTU	183.78 ± 10.99 b	0.891 ± 0.017 a	0.57 ± 0.037 abc	2.972 ± 0.170 bc	176.667 ± 7.371 b
S1	435.28 ± 106.61 a	0.966 ± 0.010 a	0.71 ± 0.037 a	4.282 ± 0.402 a	428.333 ± 106.143 a
S1TU	140.02 ± 27.82 b	0.677 ± 0.173 b	0.41 ± 0.139 cd	2.025 ± 0.692 cd	137.000 ± 26.210 b
S2	414.71 ± 97.26 a	0.905 ± 0.093 a	0.65 ± 0.102 ab	3.893 ± 0.755 ab	406.000 ± 98.190 a
S2TU	129.69 ± 43.65 b	0.620 ± 0.097 b	0.37 ± 0.036 d	1.773 ± 0.291 d	128.667 ± 44.377 b
S3	502.03 ± 15.90 a	0.976 ± 0.046 a	0.74 ± 0.183 a	4.613 ± 0.135 a	496.333 ± 14.844 a
S3TU	176.24 ± 14.98 b	0.774 ± 0.107 ab	0.47 ± 0.090 bcd	2.417 ± 0.441 cd	170.667 ± 9.866 b

Note: The letters after the numbers of peers are significant differences (p ≤ 0.05; n = 3).

). Fungal Diversity Patterns: Richness (Chao1/Observed species): Soil exhibited a significantly higher fungal richness than substrate (p < 0.05), except at the S2 stage (Substrate-S2: 72.167 vs. Soil-S2TU: 55.712). Substrate richness increased progressively with mycelial recession, while soil richness declined during S1TU-S2TU but rebounded sharply at S3TU. Diversity (Shannon/Simpson): Fungal diversity in substrate showed a sustained increase throughout recession. Soil diversity decreased during S1TU-S2TU, followed by significant recovery post-recession (S3TU). Evenness (Pielou’s index): Substrate’s evenness rose markedly from CK (0.025) to S3 (0.350), indicating disrupted evenness during mycelial growth. Soil maintained a relatively stable fungal evenness (p > 0.1), demonstrating higher environmental resilience. Bacterial Diversity Patterns (Figure 3B and

Table A2. Bacterial alpha diversity analysis table.

Sample Name	Chao1	Simpson	Pielou E	Shannon	Observed Species
CK	452.29 ± 39.14 a	0.967 ± 0.002 a	0.73 ± 0.014 a	4.455 ± 0.147 a	450.000 ± 37.723 a
CKTU	183.78 ± 10.99 b	0.891 ± 0.017 a	0.57 ± 0.037 abc	2.972 ± 0.170 bc	176.667 ± 7.371 b
S1	435.28 ± 106.61 a	0.966 ± 0.010 a	0.71 ± 0.037 a	4.282 ± 0.402 a	428.333 ± 106.143 a
S1TU	140.02 ± 27.82 b	0.677 ± 0.173 b	0.41 ± 0.139 cd	2.025 ± 0.692 cd	137.000 ± 26.210 b
S2	414.71 ± 97.26 a	0.905 ± 0.093 a	0.65 ± 0.102 ab	3.893 ± 0.755 ab	406.000 ± 98.190a
S2TU	129.69 ± 43.65 b	0.620 ± 0.097 b	0.37 ± 0.036 d	1.773 ± 0.291 d	128.667 ± 44.377 b
S3	502.03 ± 15.90 a	0.976 ± 0.046 a	0.74 ± 0.183 a	4.613 ± 0.135 a	496.333 ± 14.844 a
S3TU	176.24 ± 14.98 b	0.774 ± 0.107 ab	0.47 ± 0.090 bcd	2.417 ± 0.441 cd	170.667 ± 9.866 b

Note: The letters after the numbers of peers are significant differences (p ≤ 0.05; n = 3).

): Richness Contrast: Substrate dominated bacterial richness across all stages (Chao1/Observed species), with soil richness consistently lower except at S2/S2TU. Recession-Responsive Dynamics: Bacterial diversity declined concomitantly with active mycelial recession in both matrices, exhibiting a marginal reduction before complete recession (S3). Post-recession (S3 phase), bacterial richness showed a distinct recovery in all systems. Evenness Response: Mycelial recession induced a minimal evenness disruption in substrate bacterial communities, whereas soil bacteria experienced a substantial evenness disturbance upon complete mycelial depletion (Pielou’s index, p < 0.01). Ecological Interpretation: The inverse richness patterns between substrate and soil, coupled with recession-phase-dependent diversity fluctuations, demonstrate that Dictyophora mycelia exert inhibitory effects on co-occurring fungal/bacterial communities. This suppression was alleviated upon complete mycelial depletion.

3.4. Bata Diversity Analysis

Fungal Community Analysis (Figure 3C): Principal Coordinate Analysis (PCoA) revealed that PC1 (47.3%) and PC2 (34.5%) collectively explained 81.8% of the total variance, indicating that these two principal components adequately captured the major variation in the original data. Close spatial proximity was observed between CK and CKTU (substrate and soil with intact hyphae), as well as S3 and S3TU (substrate and soil with fully degraded hyphae), suggesting similar community compositions between these paired groups. Analysis of substrates at different hyphal regression stages demonstrated significant separation of S1, S2, and S3 from CK (PC1 axis), implying that hyphal regression in substrates likely commenced at S1. Comparative assessment of soil samples revealed a marked separation of CKTU from S1TU, S2TU, and S3TU. This spatial distribution suggests a consistent initiation timing of hyphal regression in both substrates and soils. However, soil microbial communities exhibited a greater complexity and higher stability than substrates, resulting in delayed observable regression. Bacterial Community Analysis (Figure 3D): The cumulative explanatory power of PC1 (29.4%) and PC2 (12.2%) reached 41.6%, indicating weak grouping effects among bacterial communities across hyphal regression stages. Substantial intergroup overlap further confirmed this weak partitioning. Notably, pronounced shifts in bacterial composition occurred in substrates during regression, evidenced by significant spatial separation among substrate groups. In contrast, soil samples (CKTU, S1TU, S2TU) clustered closely, demonstrating conserved bacterial community structures throughout regression.

3.5. Microbial Community Composition During Mycelial Recession

3.5.1. Fungal Community Dynamics

A total of eight substrate and soil samples across mycelial recession stages yielded 11 phyla, 29 classes, 71 orders, 141 families, and 245 genera of fungi. The dominant fungal genera with the highest relative abundance during the mycelial decline process (Figure 4A) included Phallus, Trichoderma, Scytalidium, Staphylotrichum, and Fusarium. Among these, the relative abundance of Phallus decreased as the mycelium degraded. In the substrate, the relative abundance of Phallus decreased from 99.7% to 7.0%, and in the soil, it decreased from 78.3% to 5.5%. On the other hand, Trichoderma showed an increase in relative abundance as the mycelium degraded. In the substrate, the relative abundance of Trichoderma increased from 0% to 45.2%, while in the soil, it increased from 0.1% to 55.3%.

A comparison of the changes in the dominant fungal genera in the substrate during the mycelial decline process revealed Phallus (99.7%, 61.4%, 13.0%, 7.0%) and Trichoderma (0%, 13.2%, 41.8%, 45.2%), while Scytalidium (0%, 25.2%, 39.0%, 35.6%). This indicates that Trichoderma and Scytalidium gradually became the dominant genera in the substrate as mycelial degradation progressed.

A comparison of the changes in the dominant fungal genera in the soil during the mycelial decline process revealed Phallus (78.3%, 77.5%, 94.7%, 5.5%) and Trichoderma (0.1%, 10.7%, 0.1%, 55.3%). This suggests that Trichoderma became the dominant genus in the soil, although changes in dominant genera were not noticeable in the early stages (S1TU and S2TU) when the mycelium in the soil had not yet completely degraded.

3.5.2. Bacterial Community Composition During Mycelial Recession

A total of eight substrate and soil samples across recession stages yielded 45 phyla, 141 classes, 329 orders, 540 families, and 1109 genera of bacteria. Dominant Genera Dynamics: Burkholderia–Caballeronia–Paraburkholderia, Dyella, Sphingomonas, Bacillus, and unclassified Gemmatimonadaceae constituted the predominant genera. Notably, diverse low-abundance taxa (collectively denoted as “Others”) formed a significant fraction of communities despite individual scarcity, indicating a preserved species richness throughout recession (Figure 4B).

Substrate-Specific Succession: Burkholderia–Caballeronia–Paraburkholderia (6.3%, 52%, 59%, 43%); it is a dramatic surge. Sphingomonas (0%, 0.1%, 3%, 10%); it is a progressive increase. Dyella (5.5%, 12%, 2.9%, 2.7%) shows the existence of a transient peak. Chitinophaga (17%, <0.1%, <0.1%, <0.1%) only exists in normal mycelial growth.

Soil-Specific Succession: Gemmatimonas (8.9%, 5%, 3.2%, 6.8%); fluctuating exists in the process of mycelium regression. JG30-KF-AS9 (6.4%, 6.3%, 4.7%, 4.7%) keeps stable during the process of mycelium regression. Sphingomonas (5%, 5.7%, 1.7%, 4.6%); the process of mycelium regression sees a U-shaped recovery. Dyella (0.6%, 2.4%, 4.1%, 2.7%); the phenomenon of transient increase appears in the process of mycelium decay.

D. phalloidea (Basidiomycota, genus Phallus) underwent progressive decline in abundance during mycelial recession. Concomitantly, Trichoderma emerged as the dominant successor genus in both substrate and soil matrices, suggesting its pivotal role in assembling the post-recession community. Notably, bacterial communities exhibited a U-shaped richness trajectory: initial reduction during active mycelial degradation followed by significant recovery post complete recession, as observed in both substrate and soil systems.

3.6. Microbial Community Divergence During Mycelial Recession

Fungal community heterogeneity (Figure 5A): Shared core: Four fungal genera persisted across all recession stages. Increasing β-diversity: Substrate (S3): Eleven unique genera. Soil (S3TU): Sixty-three unique genera (p < 0.001 vs. CK). Divergence magnitude: S3/S3TU > CK/S1 (Bray–Curtis dissimilarity). Bacterial community (Figure 5B): There were 56 common bacteria genera in the process of hyphae regression, but the hyphae regression led to a decrease in bacterial community species in the matrix (there were 40 endemic bacteria genera in CK but only 21 in S3), and the bacterial community species in soil were the most abundant, with a total of 138; in general, hyphae regression promoted an increase in the bacterial community in soil.

To further investigate the differences in fungal communities between substrate and soil during the mycelial regression process, LEfSe (Linear Discriminant Analysis Effect Size) analysis was performed, with an LDA score threshold of 2 (Figure 5C). This analysis identified key taxa contributing to intergroup variation. Notable shifts in community composition were observed in substrate samples S2 and S3 and in soil samples CKTU and S3TU.In the soil of the normal-growth group (CKTU), Aspergillus and Conocybe were identified as signature genera. In contrast, Mortierellales (gen. incertae sedis), Plectosphaerella, and Botryotrichum were characteristic of the completely regressed soil sample (S3TU). For substrate samples, Acremoniopsis was the only indicator genus in S2, while Burgoa, a genus within the family Hydnaceae, was uniquely enriched in S3.

Regarding bacterial community (Figure 5D) shifts during fungal mycelium recession, the soil exhibited significant enrichment of Acidobacteriota (Terriglobus, RB41) and Aneurinibacillales (Aneurinibacillales, Planifilum). Conversely, the substrate was significantly enriched in Pseudomonadales (Pseudomonas, Bordetella), Sphingomonadales (Novosphingobium, Sphingomonas), and Alicyclobacillales (Tumebacillus, Geobacillus). Notably, Nitrospira and Planifilum were predominantly enriched in soil supporting normal mycelial growth. During mycelial recession phases in the substrate, Pseudomonas was primarily enriched at stages S1 and S2, while Sphingomonas enrichment peaked at stage S3.

The changes in microbial abundance in substrate and soil flora during the process of mycelium regression (as shown in Figure 5E,F) are analyzed to further reveal the changes in microbial community in substrate and soil during the process of mycelium regression. In the order of importance of fungal communities, Fusarium is 18.7, Trichoderma is 16.6, and Columum is 16.4. The order of importance of bacterial communities: Terriglobus is 16.0, Geobacillus is 15.5, and Methylobacterium-Methylorubrum is 14.9. Among them, Fusarium is only abundant in the complete regression of soil hyphae (S3TU). The abundance of Stylosanthes increased gradually only during the change in matrix fungi community in the process of mycelium regression. Trichoderma began to fade with the mycelium, and its abundance gradually increased in matrix and soil. The thermal map in the random forest map shows that most fungi, such as Fusarium, Humicola, Penicillium, and Aspergillus, are mainly distributed in soil. However, Terriglobus and Geobacillus are only abundant in the matrix.

3.7. Niche Analysis

Figure 6A illustrates the ecological niche distribution of fungal communities in substrate and soil during the process of D. indusiatus mycelial regression. From an overall trend perspective, the median niche values in soil samples—CKTU, S1TU, S2TU, and S3TU—show a gradual decline. This may be associated with the intensity of mycelial regression or the stages of residual mycelial decomposition. In contrast, niche value distributions in substrate samples (CK, S1, S2, S3) appear relatively stable, although the median value for S3 is significantly lower than that of the other substrate groups. In terms of dispersion, niche breadth is more pronounced in soil samples. The interquartile range (IQR) for S3TU is the widest (e.g., spanning from 0.2 to 0.8), indicating intensified niche differentiation among soil fungi after mycelial regression. This could reflect increased interspecies competition or heightened resource heterogeneity. Conversely, substrate samples such as CK and S1 exhibit narrower IQRs (concentrated between 0.4 and 0.6), suggesting that environmental homogenization in the substrate has led to niche convergence among fungal taxa. Comparative analysis between groups reveals significant differences: CKTU vs. S3TU: The non-overlapping boxplots of CKTU (normal mycelial growth) and S3TU (complete mycelial regression) imply a marked shift in ecological niches, likely driven by resource depletion after complete mycelial degradation. CK vs. S3: A significant difference (p < 0.01) in median niche values between CK and S3 substrate groups suggests niche contraction, potentially resulting from Trichoderma’s dominance in the S3 substrate. S2TU: The presence of high-value outliers in the S2TU scatter plot may reflect localized, explosive growth of Phallus mycelia, temporarily expanding the ecological niches. Overall, the results of significance testing confirm that mycelial regression of D. indusiatus leads to systemic niche restructuring in both substrate and soil fungal communities.

Bacterial niche analysis (Figure 6B) revealed highly significant differences between the soil and substrate bacterial communities during mycelial recession. Bar plot results indicated higher niche breadth values in the substrate compared to soil. This suggests greater bacterial community richness within the substrate and a simpler community structure in the soil. Inter-group comparison: S1 substrate vs. S1TU soil: The niche breadth value for the S1 substrate was significantly higher than that for S1TU soil. This elevation likely corresponds to the niche occupied by enriched Pseudomonas, associated with efficient carbon utilization. S3 substrate: The niche breadth value for the S3 substrate reflects community changes occurring in the late recession phase, potentially indicating niche expansion by Sphingomonas (an aromatic compound degrader). S3TU soil: Conversely, the soil at the same stage (S3TU) maintained a low diversity, with a stable niche occupied by Nitrospira.

4. Discussion

Microscopic observation confirmed that mycelial regression in the substrate occurred significantly earlier than in the soil (consistent with reports by Wu Shaofeng et al. [6]), and microbial diversity analysis further revealed this earlier regression in the substrate compared to the soil. The stability of the fungal communities differed: both richness and evenness of soil fungal communities were significantly higher than those in the substrate, indicating that the soil microbial network possesses a stronger buffering capacity [24]. This stability delayed the mycelial regression process, likely attributable to the following: higher functional redundancy within the soil mycorrhizosphere, where saprotrophs such as Aspergillus and Conocybe can rapidly occupy vacated niches [25]; physical protection of mycelial residues by soil organic-mineral complexes, reducing degradation rates [19]. Conversely, the richness and evenness of bacterial communities were significantly higher in the substrate than in the soil, primarily due to (1) high organic matter content (e.g., lignocellulose) providing substrates for metabolically versatile bacteria like Burkholderia and Sphingomonas [26]; (2) an optimized physical structure (porosity > 40%) facilitating oxygen diffusion and microbial interactions, thereby accelerating mycelial biomass turnover [27]. In contrast, bacterial communities in the soil were constrained by nutrient limitation and spatial heterogeneity, leading to reduced diversity [28].

The microbial composition in both substrate and soil revealed the succession patterns triggered by the regression of Phallus spp. mycelia. Within fungal community shifts, the significant decline of Phallus contrasted sharply with the explosive proliferation of Trichoderma, aligning with the competitive succession model of saprotrophic fungi upon nutrient resources’ release [29]. This observation is consistent with Tan Hao et al.’s [30] findings on the impact of soil on Morchella yields. Bacterial community changes exhibited dual characteristics of the maintenance of high diversity and functional group restructuring: the dominance of the Burkholderia–Caballeronia–Paraburkholderia complex in the substrate suggests its involvement in metabolizing fungal necromass. Genes carried by this group for degrading aromatic compounds (e.g., the phn cluster) may participate in decomposing mycelial secondary metabolites [26]. The disappearance of Chitinophaga synchronized with Phallus’s mycelial regression, directly evidencing this group’s nutritional dependence on active fungal chitin [19]. The stability of Sphingomonas and Dyella underscores their role as a “core microbiome” with ecological resilience; both synthesize exopolysaccharides to withstand environmental perturbations, commonly observed in soil communities under stress [28].

Integration of LEfSe analysis and niche modeling revealed, for the first time, the mechanisms underlying microbial community differentiation between the substrate and soil during Phallus spp. mycelium recession. Fungal Community Differentiation: Soil: During the early recession stage (CKTU), the soil fungal community was dominated by Aspergillus and Conocybe. As saprophytes capable of utilizing recalcitrant organic matter (e.g., lignin), their dominance aligns with a resource conservation strategy typical of soil fungi [25]. Substrate: In the late recession stage (S3), the substrate fungal community was characterized by Burgoa (family Odontiaceae) as an indicator species. This genus likely participates in the degradation of residual mycelial chitin, and its niche expansion exhibits functional coupling with the disappearance of the bacterium Chitinophaga [31]. Bacterial Community Differentiation (LEfSe): Soil: Enrichment of Terriglobus (phylum Acidobacteriota) reflects its adaptation to oligotrophic conditions [32]. Substrate: The temporal enrichment patterns of Pseudomonas (peaking at S1–S2) and Sphingomonas (peaking at S3) correspond to their functional traits of rapid carbon source utilization and aromatic hydrocarbon degradation, respectively [33]. Microbial Interactions: Substrate, A significant collaborative degradation pathway between bacteria and fungi was identified: During recession, Pseudomonas’s degradation of labile carbon sources provided growth substrates for Trichoderma. Post-recession (S3), Sphingomonas’s degradation of aromatic metabolites (e.g., mycelial pigments) alleviated decomposition inhibition for Scytalidium [34]. Soil: Nutrient limitations led to attenuated microbial interactions. Enrichment of Acidobacteriota and Aneurinibacillales (order Aneurinibacillales) reflects oligotrophic adaptation strategies, potentially reducing nutrient availability for subsequent cultivation cycles [35].

The rapid colonization by Trichoderma, a broad-spectrum antagonist, may stem from two mechanisms: (1) Nutritional competitive advantage: Resources such as polysaccharides and chitin released by the regressing mycelia activate Trichoderma’s extracellular hydrolytic enzyme systems (e.g., chitinase Chit42) [36]. (2) Niche preemption: The “vacant niche” created by the decline of Phallus is occupied by Trichoderma, which exhibits greater environmental adaptability [18]. Furthermore, Trichoderma may act as a pathogen disrupting normal Phallus mycelial growth, significantly impacting Phallus production; species such as Trichoderma virens, Trichoderma harzianum, and Trichoderma guizhouense are major pathogens causing green mold disease in Phallus rubrovolvatus [37]. Lu Meiling’s [38] isolation and identification of pathogens from rotten P. rubrovolvatus identified Trichoderma koningii as a significantly pathogenic agent. Notably, the concurrent increase in Scytalidium in the substrate suggests functional partitioning with Trichoderma. This genus is known for its efficient cellulose degradation capability [39], and it may synergistically decompose residual cell wall polymers from the mycelia. The stability of soil fungal communities during the mid-regression stage (S1TU/S2TU), evidenced by the delayed sharp decline in Phallus abundance (94.7% → 5.5%), reflects the buffering effect of the soil microenvironment against community disruption, consistent with the “soil microbial homeostasis” theory [40].

Microbial Ecological Control Strategies for Normal D. indusiatus Mycelial Growth: Application of Antagonistic Functional Agents: Inoculating Bacillus subtilis during the mycelial recession phase significantly suppresses the over-proliferation of Trichoderma. Lipopeptide antibiotics (e.g., surfactin) secreted by B. subtilis disrupt tip growth of Trichoderma hyphae [41] while promoting colonization by D. indusiatus mycelium [42]. Precise Regulation of Substrate C/N Ratio: Adjusting the cultivation substrate’s C/N ratio (e.g., by adding nitrogen sources like soybean meal) reduces the competitive advantage of Trichoderma. A high-nitrogen environment inhibits the extracellular chitinase activity of Trichoderma [43] and enhances nutrient absorption efficiency by Phallus mycelium [18]. Soil Microbial Community Remediation: Following mycelial recession, one can amend soil with functional agents containing Sphingomonas to accelerate degradation of residual mycelium (aromatic hydrocarbon removal rate >90%), preparing the soil for subsequent D. indusiatus cultivation. Combine this with a 5% (w/w) biochar amendment to adsorb Fusarium toxins and enrich beneficial bacteria like Terriglobus [44]. Temporal Intervention of Temperature and Humidity: During the early recession phase (Stage S1), maintain substrate humidity at 60% and temperature at 28 °C to stimulate Pseudomonas’s degradation of labile carbon sources, thereby disrupting the nutrient supply to Trichoderma [45]. During the late phase (Stage S3), reduce humidity to 45% to inhibit spore dispersal (resulting in a 63% reduction in Trichoderma abundance). Additional Mitigation Strategies Against Phallus Mycelial Recession: Pre-Cultivation Site Assessment: Conduct preliminary surveys and testing of potential cultivation sites. This is crucial because Trichoderma is extensively used as a biocontrol agent in other crop production systems [46], potentially leading to high background levels. Development of Trichoderma-Resistant Varieties: Employ breeding programs for Trichoderma-resistant Phallus strains. For instance, Li et al. [47] screened Oudemansiella raphanipes and identified strains Y4, Y1, and Sgw, exhibiting high-temperature tolerance, high yields, and resistance to Trichoderma contamination.

5. Conclusions

This study systematically analyzed the succession patterns of microbial communities in both substrate and soil during the mycelial regression of Phallus spp. (D. indusiatus) through microscopic observation and high-throughput sequencing. The results revealed that mycelia underwent preferential complete regression in the substrate, while regression was delayed in the soil. A significant decline in Phallus abundance contrasted with an explosive proliferation of Trichoderma. Following regression in the soil, intensified fungal niche differentiation (indicating enhanced competition) was observed, whereas niche contraction occurred in the substrate due to the dominance of a single Trichoderma population. Within the substrate, the Burkholderia complex dominated the degradation process, whereas Sphingomonas, Gemmatimonas, and others contributed to the maintenance of stability in the soil. Bacterial diversity exhibited a rebound during the late regression stage. A primary future research direction involves determining the main drivers of D. indusiatus‘s mycelial regression to mitigate associated production losses.