Host Taxonomy and Mycelial Traits Shape Hyphosphere-Symbiotic Bacterial Communities in Cultivated Tricholomataceae Mushrooms

Host Taxonomy and Mycelial Traits Shape Hyphosphere-Symbiotic Bacterial Communities in Cultivated Tricholomataceae Mushrooms | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Host Taxonomy and Mycelial Traits Shape Hyphosphere-Symbiotic Bacterial Communities in Cultivated Tricholomataceae Mushrooms Mengzhe Gao, Shuting Zhou, Yanfei Xu, Lu Wang, Shuyu Xing, Sujing Sun This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-9530892/v1 This work is licensed under a CC BY 4.0 License Status: Under Review Version 1 posted 9 You are reading this latest preprint version Abstract Edible mushroom mycelia harbour diverse symbiotic bacteria that likely contribute to nutrient cycling, yet the factors shaping their diversity and assembly remain poorly understood. We characterized hyphal- symbiotic bacterial communities (HSBC) across eight cultivated Tricholomataceae species using 16S rRNA amplicon sequencing. Alpha diversity varied significantly among hosts, with H. marmoreus and F. filiformis_Y exhibiting the highest richness, whereas H. raphanipes showed the lowest. Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes dominated all hosts, while Tepidimicrobium, Glutamicibacter, Acinetobacter, Lactobacillus, and Sphingomonas displayed host-specific patterns. Intraspecific comparisons revealed minimal divergence between two F. filiformis strains, but the two H. marmoreus strains differed markedly in both community composition and diversity. Spearman correlation and redundancy analyses indicated that host mycelial physicochemical traits substantially influenced bacterial community differentiation. Together, these results demonstrate that host taxonomy and mycelial traits jointly shape HSBC assembly and diversity, providing a framework for understanding mushroom–bacterium interactions and promoting beneficial microbiota in cultivation. edible mushrooms hyphosphere-symbiotic bacteria host filtering microbial community assembly Tricholomataceae Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Introduction Edible mushrooms are valuable agricultural bioresources with substantial nutritional, health-promoting, and industrial value, contributing to high-quality protein provision, human health, and the development of efficient agriculture [1-3]. In recent years, shifts in dietary patterns and the growing demand for high-quality, healthy foods have driven the rapid expansion of the edible mushroom industry, which has become an important component of modern agriculture in China. In addition to being rich in proteins, amino acids, vitamins, and containing diverse bioactive compounds [4, 5]. Therefore, a better understanding of the key biological factors regulating mushroom growth, development, and metabolic traits is of considerable theoretical and practical importance for improving cultivation efficiency, yield, and product quality. In addition to conventional cultivation conditions such as temperature, pH, and substrate formulation, microbial communities, particularly hypha-associated bacteria, shape fungal growth [6, 7]. Previous studies have shown that hyphal symbiotic bacteria can profoundly shape the growth and physiological traits of their fungal hosts through nutrient transformation, metabolic interactions, synergistic extracellular enzyme activity, and the modulation of environmental adaptation. In edible mushroom cultivation systems, bacterial‒fungal interactions influence hyphal extension, substrate utilization efficiency, and fruiting body formation, underscoring the importance of these bacteria as integral components of mushroom growth and development [8]. Importantly, bacteria detected in fungal cultures are unlikely to be incidental contaminants. Increasing evidence suggests that some bacteria remain tightly associated with fungal hyphae even after isolation and repeated subculturing, notably as endohyphal bacteria or cells stably associated with hyphal surfaces [9]. For example, long-term subcultures of the ectomycorrhizal fungus Laccaria bicolor have been shown to harbor intracellular Paenibacillus spp. [10]. In the arbuscular mycorrhizal fungus Gigaspora margarita , the endobacterial symbiont Candidatus Glomeribacter gigasporarum is vertically transmitted through spores, and its elimination by continuous single-spore propagation markedly affects host spore morphology and hyphal growth [11]. Similarly, in Piriformospora indica , the endophytic bacterium Rhizobium radiobacter remains stably associated with the host under long-term axenic culture conditions, and a reduction in its abundance markedly impairs host growth, sporulation, and plant growth-promoting capacity [12]. Collectively, these findings suggest that fungus-associated bacteria are not merely incidental occupants but also active contributors to fungal life processes and ecological adaptation. Accordingly, interest in mushroom-associated microbiota has increased steadily in recent years. The bacterial communities associated with cultivated edible mushrooms, including Flammulina filiformis , Pleurotus eryngii , and Pholiota adiposa , as well as their production environments, have been increasingly investigated, revealing the complexity and dynamism of bacterial assemblages within mushroom cultivation systems [13, 14]. In addition, plate coculture experiments have shown that bacteria and their volatile metabolites can markedly affect the mycelial growth of edible mushrooms [15]. However, previous studies have focused largely on production environments, cultivation substrates, developmental stages, or individual growth-promoting strains, whereas systematic comparisons of hypha-associated bacterial communities across edible mushroom hosts remain scarce. In particular, it remains unclear whether hyphosphere symbiotic bacterial communities are structured by the host taxonomic background and whether such differentiation persists even among closely related hosts. Moreover, the contributions of host physicochemical traits and extracellular enzyme activities to bacterial community assembly and divergence have not been systematically evaluated. Here, we used eight cultivated edible mushroom species as model hosts and performed integrated analyses of hyphal growth traits, physicochemical properties, extracellular enzyme activities, and 16S rRNA gene amplicon sequencing to systematically characterize the composition and diversity patterns of these mushrooms. We further compared bacterial community variation across six genera and among different strains within species. Spearman correlation analysis together with RDA was then used to assess the contributions of the host taxonomic background and physicochemical traits to bacterial community assembly. This study aimed to elucidate edible mushroom‒bacterium interactions and the ecological principles underlying microbiome assembly, thereby providing a theoretical foundation for efficient mushroom cultivation, targeted microbiome manipulation, and the functional exploitation of mushroom‒associated bacteria Materials and Methods Fungal strains Eight commercially cultivated edible mushroom strains representing eight species were included in this study: Flammulina filiformis (two strains, including a yellow cultivar), Hypsizygus marmoreus (two strains, including a brown cultivar), Lyophyllum decastes , Hymenopellis raphanipes , Pleurotus giganteus and Macrocybe gigantea . All the strains were isolated from commercially cultivated fruiting bodies. Pure cultures were established via tissue isolation and maintained on potato dextrose agar (PDA) (Supplementary Fig. 1). Detailed information on the strain origin is provided in Supplementary Table 1. Culture media An enriched potato dextrose agar (PDA) medium was prepared from potato infusions obtained from 200 g fresh potato and supplemented with 20 g glucose, 3 g yeast extract, 3 g peptone, 1.5 g KH 2 PO 4 and 1.5 g MgSO 4 per liter. For the solid medium, 20 g agar was added per liter. The LBL medium was prepared from potato infusions obtained from 200 g fresh potato and supplemented with 20 g lactose, 2 g NH 4 NO 3 , 0.5 g MgSO 4 , 1.5 g KH 2 PO 4 and 0.08 g of bromothymol blue (BTB) per liter; the pH was adjusted to 7.0. The media were autoclaved at 121 °C for 20 min. All the strains were cultured at 25 °C in the dark. ITS amplification and phylogenetic analysis The fruiting bodies were cleaned and surface-sterilized, and 5 mm thick tissue blocks were aseptically placed on PDA. The cultures were incubated at 25 °C, and the emerging hyphae were subcultured to obtain pure isolates. Genomic DNA was extracted from fresh mycelia via a commercial kit. The ITS region was amplified via the universal fungal primers ITS4 and ITS5 [16]. After verification by agarose gel electrophoresis, the PCR products were sent for commercial sequencing. Sequences with high-quality chromatograms were aligned via ClustalW [17], and low-quality terminal bases were trimmed. Phylogenetic analysis was performed in MEGA 11.0. A neighbor‒joining tree was constructed via the Kimura two-parameter model, and branch support was assessed with 1,000 bootstrap replicates. The resulting ITS phylogeny was used to infer the taxonomic identities and phylogenetic relationships of the eight strains. Mycelial growth and physicochemical measurements Strains were cultured on PDA plates at 25 °C in the dark, and the mycelial growth rate (mm day⁻¹) was measured as previously described [18, 19]. Mycelial biomass was determined from liquid cultures. After cultivation, the mycelia were harvested, dried to a constant weight and weighed. Biomass was calculated as follows: Biomass (g 100 ml⁻¹) = mycelium dry weight (g)/culture volume (ml) × 100 [20]. The pH of the fermentation broth was measured via a pH meter. Mycelium total carbon content was determined according to T/NAIA 070-2021 via the potassium dichromate titration method [21]. Mycelium total nitrogen content was determined via the Kjeldahl method with an automatic Kjeldahl nitrogen analyzer according to NY/T 1121.24-2012 [22, 23]. Laccase activity was assayed via the ABTS radical cation decolorization method with 2,2′-azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) as the substrate [24]. The crude polysaccharide and soluble protein contents, as well as amylase, cellulase and peroxidase activities, were determined via commercial kits from Enzyme-linked Biotechnology Co., Ltd., according to the manufacturer’s instructions. DNA extraction and 16S rRNA gene sequencing Total DNA was extracted from mycelial samples via a commercial DNA extraction kit [25]. The V3-V4 region of the bacterial 16S rRNA gene was amplified via the universal primer pair 338F (5′-ACTCCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). Each PCR mixture (20 μL) contained 4 μL of 5 × TransStart FastPfu buffer, 2 μL of 2.5 mM dNTPs, 0.8 μL of each primer (5 μM), 0.4 μL of TransStart FastPfu DNA polymerase, 10 ng of template DNA, and nuclease-free water to 20 μL [26]. PCR was performed on a GeneAmp 9700 thermocycler (ABI) under the following conditions: 95 °C for 3 min; 27 cycles of 95 °C for 30 s, 55 °C for 30 s and 72 °C for 30 s; and a final extension at 72 °C for 10 min. Amplicons were sequenced on an Illumina MiSeq PE300 platform [27, 28]. Sequence processing and community analysis Raw reads were merged via FLASH [29], and adapter and primer sequences were removed via Cutadapt [30] to obtain high-quality reads. Operational taxonomic units (OTUs) were clustered at 97% sequence similarity, and representative sequences were taxonomically assigned via the SILVA database [31]. Low-abundance OTUs, as well as OTUs assigned to chloroplasts or mitochondria, were removed. The samples were then rarefied to the minimum sequencing depth before downstream analyses. On the basis of the rarefied OTU table, the Chao1, ACE, Shannon, and Simpson indices were calculated to assess alpha diversity. Beta diversity was evaluated via principal coordinate analysis (PCoA), non-metric (NMDS) and permutational multivariate analysis of variance (PERMANOVA) on the basis of Bray‒Curtis distances. The top 10 dominant bacterial phyla and the top 20 dominant bacterial genera were used to characterize the community composition. Differentially abundant taxa among groups were identified via LEfSe (LDA=2, p <0.05). Statistical analysis Statistical analyses were performed using GraphPad Prism. Differences among groups were assessed by one-way analysis of variance (ANOVA), followed by appropriate post hoc tests. Community analyses and visualizations were performed via the Majorbio Cloud Platform (https://www.majorbio.com/). The graphical abstracts were created via Adobe Illustrator 2021. Results ITS-based phylogeny and taxonomic relationships of the fungal strains Fruiting body morphology, together with mycelial growth and colony characteristics on PDA-enriched media, is shown for all the tested strains in Supplementary Fig. 1. ITS-based phylogenetic analysis using reference sequences from GenBank was performed to infer the relationships among the tested strains (Supplementary Fig. 2a). The eight strains were resolved into six phylogenetic clades corresponding to Flammulina (two strains), Hypsizygus (two strains), Hymenopellis (one strain), Lyophyllum (one strain), Pleurotus (one strain), and Macrocybe (one strain). This classification was consistent with the observed fruiting body morphology. The yellow and white F. filiformis isolates, as well as the white and brown H. marmoreus isolates, represented different strains of the same species. Variation in host mycelial physicochemical traits The nutritional and physiological traits of the eight edible mushroom mycelia are shown in Fig. 1a-i. F. filiformis _Y presented the highest mycelial growth rate (11.62 mm day⁻¹), followed by F. filiformis _W (10.01 mm day⁻¹) and H. raphanipes (9.41 mm day⁻¹) (Supplementary Fig. 2b). Mycelial biomass in liquid culture showed an aimilar pattern, indicating a close association between biomass accumulation and mycelial growth (Supplementary Fig. 2c). Among the tested strains, F. filiformis _W, H. marmoreus _W, and H. marmoreus _B presented relatively high pH values, whereas M. gigantea presented the lowest pH value (Supplementary Fig. 2d). As shown in Fig. 1a-c, significant differences in total carbon content, total nitrogen content, and the carbon-to-nitrogen ratio were observed among the tested strains. F. filiformis _Y presented the highest total carbon content, whereas L. decastes presented the lowest value. The total nitrogen content varied within a relatively narrow range, with M. gigantea , F. filiformis_W , and H. marmoreus _B showing relatively high values, and F. filiformis _Y and P. giganteus presented relatively low values. For the carbon-to-nitrogen ratio, F. filiformis _Y presented the highest value, whereas H. marmoreus _B, L. decastes , and M. gigantea presented relatively low values. The crude polysaccharide content was highest in M. gigantea (166.65 mg g⁻¹), followed by H. marmoreus _B (94.72 mg g⁻¹) (Fig. 1d). The soluble protein content ranged from 5.41 to 7.61 mg g⁻¹, with the highest value in H. marmoreus _B and the lowest in L. decastes (Fig. 1e). F. filiformis _Y presented the highest amylase activity, followed by F. filiformis -W and H. raphanipes , whereas H. marmoreus _Y and H. marmoreus _W presented the lowest activities. (Fig. 1f). The highest cellulase activity was observed in H. marmoreus _W (11.47 U), whereas the lowest activities were detected in H. marmoreus _B and M. gigantea , at 3.20 and 2.40 U, respectively (Fig. 1g). In contrast, a similar trend was observed for laccase and peroxidase activities: both activities were relatively high in L.decastes , F. filiformis _Y, F. filiformis _W, and H. raphanipes but low in H. marmoreus _W, H. marmoreus _B, and M. gigantea (Fig. 1h-i). HSBC differed among the six host genera High-throughput sequencing generated 1,523,460 valid reads, which were clustered into 1,028 OTUs. These OTUs were assigned to 25 phyla, 49 classes, 121 orders, 230 families and 527 genera (Supplementary Table 2). Venn analysis revealed that 396 OTUs were shared among the six host genera, whereas Flammulina , Hymenopellis , Hypsizygus , Lyophyllum , Pleurotus , and Macrocybe harbored 2, 4, 23, 7, 2 and 12 unique OTUs (Fig. 3a), respectively. Alpha-diversity analysis revealed that HSBC associated with H. marmoreus and F. filiformis _Y presented relatively high richness, whereas H. raphanipes exhibited the lowest alpha diversity (Fig. 2a-b). The Shannon and Simpson indices suggested broadly similar evenness patterns among the host genera, although Hypsizygus and Pleurotus were more distinct from the other groups (Fig. 2b-c). At the OTU level, PCA and NMDS showed tight clustering within groups, indicating good reproducibility, although a few P. giganteus and M. gigantea samples deviated from their group centroids (Fig. 3b-c). At the phylum level, all six host genera were dominated by Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes. Proteobacteria was particularly abundant in L. decastes (47.15%) and P. giganteus (44.80%), whereas Firmicutes was most abundant in H. marmoreus (43.60%) (Fig. 3d). At the genus level, Tepidimicrobium was the dominant genus across all six host genera. Glutamicibacter was enriched in M. gigantea , H. marmoreus and F. filiformis ; Acinetobacter was abundant in L. decastes and H. marmoreus ; Lactobacillus was significantly enriched in H. marmoreus ; and Sphingomonas was most abundant in H. raphanipes (Fig. 3e-f). LEfSe further identified distinct biomarker taxa among host genera, indicating clear host-dependent selection on symbiotic bacterial communities (Fig. 3g). Intraspecific variation differed between F. filiformis and Hypsizygus marmoreus To further assess intraspecific variation, two F. filiformis strains and two H. marmoreus strains were compared. Venn analysis revealed that the two F. filiformis strains shared 444 genera, with 19 and 35 unique genera in the yellow and white strains, respectively (Fig. 4a). The two H. marmoreus strains shared 497 genera, with 46 and 19 unique genera among the white and brown strains, respectively (Fig. 4b). No significant differences in alpha diversity were observed between the two F. filiformis strains (Fig. 4e-h). NMDS further revealed that the hyphosphere bacterial community structure did not differ significantly between the two strains. At the genus and phylum levels, none of the 20 most abundant genera differed significantly between the two strains (Fig. 5a-d). In contrast, the two H. marmoreus strains differed markedly. Compared with the brown strain, the white strain presented significantly greater alpha diversity indices (Fig. 4i-k). NMDS clearly separated the two groups (Fig. 4c). At the phylum level, no significant differences were observed between the two H. marmoreus strains (Fig. 5e-f), At the genus level, six taxa, Tepidimicrobium , Lactobacillus , Glutamicibacter , Ruminococcaceae_UCG-014, Ralstonia and Erysipelothrix , differed significantly between the two strains (Fig. 5g-h). Physicochemical traits are associated with bacterial community structure Spearman correlation analysis revealed distinct associations between host mycelial physicochemical traits and bacterial genera. Hierarchical clustering revealed that TN and TS grouped together, C/N clustered closely with SP, and Amy clustered most closely with Lac, which was further grouped with POD and Cel (Fig. 6). These results suggest that these variables exerted broadly similar effects on genus-level distribution patterns. Several genera, including Tepidimicrobium , Fermentimonas , Proteiniphilum , Amphibacillus , Erysipelothrix , Actinomyces , PMMR1, Ralstonia , and Pelomonas , were positively correlated with TC, C/N, and SP but negatively correlated with POD, Amy, and Lac. In contrast, Bacillus , Brevundimonas , and Acinetobacter were relatively strongly positively correlated with Cel and POD but negatively correlated with C/N. Enterobacter had the strongest positive correlation with SP and was also positively correlated with C/N and Lac but negatively correlated with TN. In addition, Glutamicibacter was positively correlated with TS, Ochrobactrum was negatively correlatedwith TN, and Methylobacterium was negatively correlated with C/N. Overall, genus-specific associations with host physicochemical traits were evident, with Enterobacter showing the strongest response. RDA revealed that the first two axes explained 21.64% and 11.13% of the total variation, respectively, accounting for 32.77% of the overall variation. TS, TN and POD were associated mainly with the negative side of RDA1, whereas SP, C/N and Lac were associated with the positive side of RDA1; Amy and TC were associated primarily with the negative side of RDA2. M. gigantea was positioned on the positive side of RDA1, which is consistent with SP, C/N and Lac, whereas H. marmoreus was distributed in the positive RDA1 and negative RDA2 spaces in association with Lac, Amy and TC. In contrast, F. filiformis and P. giganteus were located mainly on the negative side of RDA1, closer to TS, TN and POD. H. raphanipes clustered near the origin, indicating weaker relationships with the measured physicochemical variables, whereas L. decastes occupied the negative RDA1 and positive RDA2 space and was separated from most of the environmental vectors (Fig. 7). Discussion Host taxonomic background drives the differentiation of HSBC In the present study, HSBC differed significantly among the host genera, although all the groups were consistently dominated by Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes. This pattern suggests that hyphosphere microbiome assembly is not entirely stochastic but instead reflects the combined effects of core conservation and host specificity within a shared environmental bacterial pool. Similar patterns have also been observed in other studies of fungus-associated microbiomes. Liu et al. [13] used high-throughput 16S rRNA amplicon sequencing to characterize microbial communities associated with F. filiformis from different production regions and reported a conserved set of dominant taxa, whereas the relative abundances of Pseudomonas and Lactobacillus varied markedly among locations. Similarly, Chen et al. [14] reported that the bacterial community associated with Pleurotus eryngii changed significantly across developmental stages and that some bacterial taxa were positively correlated with the mycelial growth rate, indicating their potential growth-promoting effects. Together, Collectively, these findings indicate that the mushroom-associated microbiota exhibits both stability and ecological plasticity, with bacterial–fungal associations being widespread across phylogenetically diverse fungi rather than exceptional occurrences[32]. Similarly, extraradical hyphae of arbuscular mycorrhizal fungi have been shown to recruit conserved and reproducible bacterial communities, suggesting that fungal hyphae can themselves actively shape the surrounding microbial microenvironment [33]. These observations imply that the host taxonomic background in this context reflects more than differences in identity or nomenclature alone and may represent underlying differences in the host resource supply, metabolic strategy, and ecological niche, all of which can contribute to the differentiation of HSBC. Relationships between microbiome divergence and host trait variation among closely related hosts To further evaluate the effects of host phylogenetic relatedness and host trait variation on the hyphosphere microbiota, we compared the bacterial communities associated with two strains of F. filiformis and two strains of H. marmoreus . The results revealed that neither the community structure nor the predicted functions differed significantly between the two F. filiformis strains. In contrast, the community compositions of the two H. marmoreus strains differed significantly, and alpha diversity analysis indicated that phylogenetic proximity does not necessarily result in microbiome convergence. Instead, when the host taxonomic background is relatively constant, differences in host phenotype and metabolism may become important drivers of microbiome assembly. This interpretation is consistent with previous studies [34]. reported that bacterial communities associated with different fungal phylogenetic groups differed significantly and that these differences could be partly explained by host chemical traits, particularly pH and the C/N ratio. These results suggest that both the host phylogenetic background and the host physicochemical characteristics jointly shape the assembly of fungus-associated microbiota. Emmett et al. [33] further demonstrated that the bacterial communities associated with the extraradical hyphae of arbuscular mycorrhizal fungi remained highly conserved across different soil backgrounds and nutrient conditions. This finding suggests that stable and long-term interactions may exist between fungal hosts and hypha-associated or hypha-internal microbiota and that coadaptation, or even coevolution, may occur in certain systems. In the present study, the contrasting patterns observed between the Flammulina and Hypsizygus strains may reflect the hierarchical interplay between phylogenetic constraints and host trait divergence. When key physicochemical traits differ only slightly among closely related hosts, phylogenetic constraints may remain the dominant factor maintaining microbiome conservatism. However, when host physicochemical differences become more pronounced, ecological niche differentiation and the associated environmental filtering may override the effect of phylogenetic similarity, thereby promoting shifts in both community composition and potential functional profiles. Host regulation of hyphosphere bacterial community assembly Further Spearman correlation analysis revealed that host physicochemical traits, including total carbon, crude polysaccharide, soluble protein, pH, and the C/N ratio, were significantly associated with the distribution of dominant bacterial genera and overall community structure. These results suggest that host traits may shape community assembly through both resource supply and environmental filtering. The total carbon, crude polysaccharide, and soluble protein contents were positively correlated with several dominant genera, indicating that increased availability of carbon and nitrogen resources may facilitate the colonization and enrichment of specific bacterial taxa. In contrast, the C/N ratio and the activities of some extracellular enzymes were negatively correlated with those of certain genera, suggesting that shifts in host nutritional conditions and metabolic status may restrict the proliferation of particular bacterial taxa and thereby reshape community composition. Previous hyphosphere studies have shown that carbon-rich compounds released from fungal hyphae can significantly alter the composition and functional potential of surrounding microbial communities, thereby making the hyphosphere a distinct ecological niche relative to the bulk environment [35]. In addition, phosphorus forms, organic phosphorus levels, and hyphal continuity have all been shown to influence the assembly of HSBC [36]. Studies of soil microbial communities have likewise identified pH as a major environmental driver of bacterial community differentiation [37]. Therefore, the coexistence of positive and negative correlations between host physicochemical traits and bacterial taxa suggests that host mycelia may selectively filter hyphosphere bacteria through variations in nutrient inputs, microenvironmental conditions, and substrate transformation. The RDA results provided further support for this interpretation. SP, C/N, and Lac presented similar vector orientations, whereas TC and Amy presented broadly consistent distribution patterns, suggesting that these variables may jointly reflect the resource status and metabolic activity surrounding host hyphae and exert coordinated effects on the bacterial community structure. In contrast, TN, TS, and POD were oriented toward the left side of the ordination space. Notably, the Cel vector was short and close to the origin, indicating a limited contribution to the first two axes and suggesting that cellulose-related processes were unlikely to be the primary drivers of community divergence in this system. This interpretation is consistent with previous studies showing that microbial community composition is closely associated with specific enzyme activities, carbon chemistry, and resource availability and that differences in substrate utilization between bacteria and fungi can further promote niche differentiation and community reorganization [38, 39]. In addition, different hosts appeared to exert varying strengths of selection on hypha-associated bacterial communities. The P. giganteus group was distributed mainly on the right side of the ordination space and was closely aligned with SP, C/N, TC, Amy, and Lac, suggesting a relatively stable association between community structure and these physicochemical traits. In contrast, the L. decastes group was distributed mainly in the upper-left region and was clearly separated from P. giganteus , indicating that different host groups may possess distinct nutrient-use strategies or metabolic characteristics that promote directional differentiation of bacterial communities. Previous studies have shown that bacteria associated with edible mushrooms not only contribute to nutrient supply, mycelial growth, and fruiting body formation but also vary significantly with host type and local environmental conditions [40, 41]. Taken together, these findings suggest that HSBC assembly is jointly driven by the host taxonomic background and host physicochemical traits. Conclusion Our results show that host taxonomic identity underlies the broad differentiation of HSBC across edible mushroom genera, giving rise to microbial assemblages that contain both conserved core taxa and host-specific components. Among closely related hosts, mycelial nutritional and physicochemical traits have emerged as increasingly important drivers of bacterial community reassembly. Together, these findings indicate that hyphosphere microbiome assembly is jointly shaped by host evolutionary history and host-associated environmental factors. This work provides insight into the hierarchical contributions of host taxonomy and host traits to microbiome assembly and offers a theoretical framework for the targeted regulation of beneficial microbiota in edible mushroom cultivation. Declarations Acknowledgments This work was supported by the Project of Modern Agricultural Industrial Technology System of Edible Fungi in Fujian Province. Author Contributions All the authors contributed to the study's conception and design. Mengzhe Gao, Yanfei Xu and Lu Wang; investigation: Mengzhe Gao and Shuting Zhou; writing—original draft preparation: Shuyu Xing; writing—review and editing: Sujing Sun: Conceptualization, supervision, resources, funding acquisition, writing—review and editing. All the authors commented on previous versions of the manuscript and approved its final version. Funding Data availability All the data generated or analyzed during this study are included in this published article and its supplementary information files. Conflict of interest The authors declare that they have no competing interests. Open Access References Valverde ME, Hernández-Pérez T, Paredes-López O (2015) Edible mushrooms: improving human health and promoting quality life. 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Nucleic Acids Res 41:D590-596. https://doi.org/10.1093/nar/gks1219 Robinson AJ, House GL, Morales DP, Kelliher JM, Gallegos-Graves V, LeBrun ES, Davenport KW, Palmieri F, Lohberger A, Bregnard D, Estoppey A, Buffi M, Paul C, Junier T, Hervé V, Cailleau G, Lupini S, Nguyen HN, Zheng AO, Gimenes LJ, Bindschedller S, Rodrigues DF, Werner JH, Young JD, Junier P, Chain PSG (2021) Widespread bacterial diversity within the bacteriome of fungi. Commun Biol 4:1168. https://doi.org/10.1038/s42003-021-02693-y Emmett BD, Lévesque-Tremblay V, Harrison MJ (2021) Conserved and reproducible bacterial communities associate with extraradical hyphae of arbuscular mycorrhizal fungi. Isme j 15:2276-2288. https://doi.org/10.1038/s41396-021-00920-2 Pent M, Bahram M, Põldmaa K (2020) Fruitbody chemistry underlies the structure of endofungal bacterial communities across fungal guilds and phylogenetic groups. Isme j 14:2131-2141. https://doi.org/10.1038/s41396-020-0674-7 Zhang L, Zhou J, George TS, Limpens E, Feng G (2022) Arbuscular mycorrhizal fungi conducting the hyphosphere bacterial orchestra. Trends Plant Sci 27:402-411. https://doi.org/10.1016/j.tplants.2021.10.008 Wang F, Kertesz MA, Feng G (2019) Phosphorus forms affect the hyphosphere bacterial community involved in soil organic phosphorus turnover. Mycorrhiza 29:351-362. https://doi.org/10.1007/s00572-019-00896-0 Rousk J, Bååth E, Brookes PC, Lauber CL, Lozupone C, Caporaso JG, Knight R, Fierer N (2010) Soil bacterial and fungal communities across a pH gradient in an arable soil. Isme j 4:1340-1351. https://doi.org/10.1038/ismej.2010.58 Li Y, Nie C, Liu Y, Du W, He P (2019) Soil microbial community composition closely associates with specific enzyme activities and soil carbon chemistry in a long-term nitrogen fertilized grassland. Science of The Total Environment 654:264-274. https://doi.org/https://doi.org/10.1016/j.scitotenv.2018.11.031 Wang C, Kuzyakov Y (2024) Mechanisms and implications of bacterial–fungal competition for soil resources. The ISME Journal 18:wrae073. https://doi.org/10.1093/ismejo/wrae073 Pent M, Põldmaa K, Bahram M (2017) Bacterial communities in boreal forest mushrooms are shaped both by soil parameters and host identity. Front Microbiol 8:836. https://doi.org/10.3389/fmicb.2017.00836 Sun K, Jiang HJ, Pan YT, Lu F, Zhu Q, Ma CY, Zhang AY, Zhou JY, Zhang W, Dai CC (2023) Hyphosphere microorganisms facilitate hyphal spreading and root colonization of plant symbiotic fungus in ammonium-enriched soil. Isme j 17:1626-1638. https://doi.org/10.1038/s41396-023-01476-z Additional Declarations No competing interests reported. Supplementary Files Supplementarytable.docx supplementaryFig.1.tif supplementaryFig.2.tif Cite Share Download PDF Status: Under Review Version 1 posted Reviewers agreed at journal 04 May, 2026 Reviewers agreed at journal 04 May, 2026 Reviewers agreed at journal 03 May, 2026 Reviewers agreed at journal 01 May, 2026 Reviewers agreed at journal 29 Apr, 2026 Reviewers invited by journal 29 Apr, 2026 Editor assigned by journal 29 Apr, 2026 Submission checks completed at journal 29 Apr, 2026 First submitted to journal 26 Apr, 2026 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-9530892","acceptedTermsAndConditions":true,"allowDirectSubmit":false,"archivedVersions":[],"articleType":"Research Article","associatedPublications":[],"authors":[{"id":634247571,"identity":"e8872ca7-3067-4504-858f-c1d2031bb691","order_by":0,"name":"Mengzhe Gao","email":"","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":false,"prefix":"","firstName":"Mengzhe","middleName":"","lastName":"Gao","suffix":""},{"id":634247574,"identity":"5f2334c9-4427-4a5d-a4c4-f815f19c4967","order_by":1,"name":"Shuting Zhou","email":"","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":false,"prefix":"","firstName":"Shuting","middleName":"","lastName":"Zhou","suffix":""},{"id":634247575,"identity":"648f3b11-9fef-49cb-9ff8-0777f651b299","order_by":2,"name":"Yanfei Xu","email":"","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":false,"prefix":"","firstName":"Yanfei","middleName":"","lastName":"Xu","suffix":""},{"id":634247580,"identity":"a78e491e-f898-46a5-af58-63c9d9c4b6e9","order_by":3,"name":"Lu Wang","email":"","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":false,"prefix":"","firstName":"Lu","middleName":"","lastName":"Wang","suffix":""},{"id":634247584,"identity":"21bd1161-255c-4480-9cba-63a0a126dfc0","order_by":4,"name":"Shuyu Xing","email":"","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":false,"prefix":"","firstName":"Shuyu","middleName":"","lastName":"Xing","suffix":""},{"id":634247586,"identity":"a7aadd47-175e-480f-9b9f-d38c2382cb1e","order_by":5,"name":"Sujing Sun","email":"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAA3klEQVRIie3RsYrCMBzH8f8heEtDN0kpmFdI6erD/IPQ6R7AsXCSZ6iI75DR8V8KneJe8OAEVw8KXQXPji4mo2A+Q6bfFxICEASviAPez0U0hYaoX/knxTz+aFVdWb9k1OTJt80btvYoxLY8D2w/Uaa1PbESRDyj54n8oSJldqqMPRhK9pBttuhIOBYTpiNlunuSWUB5dCSiwuXANFfm93IipT0S6BBTpmWelBao9klkh0Wy0ziPoZV1abn7LaL6WvZ/+jZ+5Xm4rhYiTl0Xg+hhwV3z0Sf5rIIgCN7ZP1xmTtFinJ1bAAAAAElFTkSuQmCC","orcid":"","institution":"Fujian Agriculture and Forestry University","correspondingAuthor":true,"prefix":"","firstName":"Sujing","middleName":"","lastName":"Sun","suffix":""}],"badges":[],"createdAt":"2026-04-26 09:40:15","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-9530892/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-9530892/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":108808214,"identity":"784d6da4-1b62-4340-b53c-32426233fe15","added_by":"auto","created_at":"2026-05-08 15:40:31","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":16053806,"visible":true,"origin":"","legend":"\u003cp\u003eHost mycelial physicochemical traits of eight cultivated edible mushroom strains. (a) Total carbon content. (b) Total nitrogen content. (c) Carbon-to-nitrogen ratio. (d) Total sugar. (e) Soluble protein content. (f) Amylase activity. (g) Cellulase activity. (h) Laccase activity. (i) Peroxidase activity. The data are presented as the means ± standard errors. Different letters above the bars indicate significant differences among strains (one-way ANOVA followed by post hoc tests, \u003cem\u003ep\u003c/em\u003e \u0026lt; 0.05).\u003c/p\u003e","description":"","filename":"fig.1.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/7551a80299f8de6707c824ca.png"},{"id":108808171,"identity":"605084ff-3cba-46c1-b374-e0a83a0d6152","added_by":"auto","created_at":"2026-05-08 15:40:13","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":2530407,"visible":true,"origin":"","legend":"\u003cp\u003eAlpha diversity indices of the HSBC among the six host genera of edible mushrooms. (a) Chao1. (b) ACE. (c) Simpson. (d) Shannon. Different letters indicate significant differences among host genera (p \u0026lt; 0.05).\u003c/p\u003e","description":"","filename":"Fig.2.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/3cab8f7f2ae0577e9e40c4c7.png"},{"id":108810054,"identity":"7f10ed59-a0d2-4dea-8d53-7ae8b4c9c6c3","added_by":"auto","created_at":"2026-05-08 15:57:16","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":17887717,"visible":true,"origin":"","legend":"\u003cp\u003eComposition, beta diversity, and differential taxa of HSBC among the six host genera of edible mushrooms. (A) Venn diagram showing the numbers of shared and unique OTUs among \u003cem\u003eFlammulina\u003c/em\u003e, \u003cem\u003eHymenopellis\u003c/em\u003e, \u003cem\u003eHypsizygus\u003c/em\u003e, \u003cem\u003eLyophyllum\u003c/em\u003e, \u003cem\u003ePleurotus\u003c/em\u003e, and \u003cem\u003eMacrocybe\u003c/em\u003e. (B) Principal coordinate analysis (PCoA) based on Bray–Curtis distances showing differences in bacterial community structure among the six host genera. (C) Non-metric (NMDS) analysis of the bacterial community composition among the six host genera. (D) Relative abundances of the dominant bacterial phyla across the six host genera. (E) Relative abundances of the dominant bacterial genera across the six host genera. (F) LEfSe analysis showing bacterial taxa with significantly enriched differences among host genera (LDA score \u0026gt; 2.0,\u003cem\u003e p\u003c/em\u003e \u0026lt; 0.05). (G) Cladogram of the differentially abundant taxa identified via LEfSe.\u003c/p\u003e","description":"","filename":"Fig.3.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/9273d76fa1d455e52a1d0705.png"},{"id":108808215,"identity":"1a5f5831-4a73-4b7c-bb2f-a2db6cd1b484","added_by":"auto","created_at":"2026-05-08 15:40:31","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":8223791,"visible":true,"origin":"","legend":"\u003cp\u003eIntraspecific variation in hyphosphere symbiotic bacterial communities between two \u003cem\u003eFlammulina filiformis\u003c/em\u003e strains and two \u003cem\u003eHypsizygus marmoreus\u003c/em\u003e strains. (A) Venn diagram of bacterial genera shared by and unique to the yellow and white strains of \u003cem\u003eF. filiformis\u003c/em\u003e. (B) Venn diagram of bacterial genera that are shared and unique between the white and brown strains of \u003cem\u003eH. marmoreus\u003c/em\u003e. (C) NMDS ordination of the bacterial community structure of the two\u003cem\u003e F. filiformis\u003c/em\u003estrains. (D) NMDS ordination of the bacterial community structure in the two \u003cem\u003eH. marmoreus \u003c/em\u003estrains. (E–H) Alpha diversity indices (ACE, Chao1, Simpson, and Shannon indices) of the two \u003cem\u003eF. filiformis\u003c/em\u003e strains. (I–L) Alpha diversity indices (ACE, Chao1, Simpson, and Shannon indices) of the two \u003cem\u003eH. marmoreus\u003c/em\u003estrains.\u003c/p\u003e","description":"","filename":"Fig.4.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/88b753b73feee4cef6454952.png"},{"id":108809880,"identity":"1a0b0271-80e7-43b9-b2d1-4340d9807cbc","added_by":"auto","created_at":"2026-05-08 15:56:03","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":7056794,"visible":true,"origin":"","legend":"\u003cp\u003eComparison of the HSBC structure between two \u003cem\u003eFlammulina filiformis\u003c/em\u003e strains and two \u003cem\u003eHypsizygus marmoreus\u003c/em\u003e strains. (A, B) Dominant bacterial phyla and their differential abundance between the yellow and white strains of \u003cem\u003eF. filiformis\u003c/em\u003e. (C, D) Dominant bacterial genera and their differential abundance between the two \u003cem\u003eF. filiformis\u003c/em\u003e strains. (E, F) Dominant bacterial phyla and their differential abundance between the white and brown strains of \u003cem\u003eH. marmoreus\u003c/em\u003e. (G, H) Dominant bacterial genera and their differential abundance between the two \u003cem\u003eH. marmoreus\u003c/em\u003e strains.\u003c/p\u003e","description":"","filename":"Fig.5.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/9afc65360bd54724641a20a1.png"},{"id":108808021,"identity":"c67eb1d4-9d2c-436f-b88a-2d47444292b2","added_by":"auto","created_at":"2026-05-08 15:39:03","extension":"png","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":2214117,"visible":true,"origin":"","legend":"\u003cp\u003eSpearman correlation heatmap showing the relationships between dominant bacterial genera and host mycelial physicochemical traits. The color gradients indicate the strength and direction of the correlations in the panel.\u003c/p\u003e","description":"","filename":"Fig.6.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/f5f02d6fc5a73cfbadd505ec.png"},{"id":108808207,"identity":"8d57ca15-ec74-47ac-b0c7-25c2f02e9920","added_by":"auto","created_at":"2026-05-08 15:40:26","extension":"png","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":3117942,"visible":true,"origin":"","legend":"\u003cp\u003eRedundancy analysis (RDA) showing the relationships between host genera and physicochemical variables. The arrows indicate explanatory variables in the panel.\u003c/p\u003e","description":"","filename":"Fig.7.png","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/246422a20e235d83c54e665e.png"},{"id":108815212,"identity":"a2f7d341-7dff-4f2e-9094-061ec38f34a9","added_by":"auto","created_at":"2026-05-08 16:21:53","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":44323613,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/06fd88fa-bede-41f0-ae78-6e46ed3f72d2.pdf"},{"id":108810179,"identity":"e704c8f6-ab47-4117-9191-1fd31808cd05","added_by":"auto","created_at":"2026-05-08 15:57:47","extension":"docx","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":16582,"visible":true,"origin":"","legend":"","description":"","filename":"Supplementarytable.docx","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/4eb13c35a6a4e3304aba6b58.docx"},{"id":108808206,"identity":"e07225c8-040d-4391-84b0-2c76211abeb7","added_by":"auto","created_at":"2026-05-08 15:40:26","extension":"tif","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":54221816,"visible":true,"origin":"","legend":"","description":"","filename":"supplementaryFig.1.tif","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/2f08892819eae5c6d66c347f.tif"},{"id":108808208,"identity":"03bbba3c-e9eb-4523-9650-af561cc3bd97","added_by":"auto","created_at":"2026-05-08 15:40:26","extension":"tif","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":49261612,"visible":true,"origin":"","legend":"","description":"","filename":"supplementaryFig.2.tif","url":"https://assets-eu.researchsquare.com/files/rs-9530892/v1/4221772917dea82c7ae70145.tif"}],"financialInterests":"No competing interests reported.","formattedTitle":"Host Taxonomy and Mycelial Traits Shape Hyphosphere-Symbiotic Bacterial Communities in Cultivated Tricholomataceae Mushrooms","fulltext":[{"header":"Introduction","content":"\u003cp\u003eEdible mushrooms are valuable agricultural bioresources with substantial nutritional, health-promoting, and industrial value, contributing to high-quality protein provision, human health, and the development of efficient agriculture [1-3]. In recent years, shifts in dietary patterns and the growing demand for high-quality, healthy foods have driven the rapid expansion of the edible mushroom industry, which has become an important component of modern agriculture in China. In addition to being rich in proteins, amino acids, vitamins, and containing diverse bioactive compounds [4, 5]. Therefore, a better understanding of the key biological factors regulating mushroom growth, development, and metabolic traits is of considerable theoretical and practical importance for improving cultivation efficiency, yield, and product quality. In addition to conventional cultivation conditions such as temperature, pH, and substrate formulation, microbial communities, particularly hypha-associated bacteria, shape fungal growth [6, 7]. Previous studies have shown that hyphal symbiotic bacteria can profoundly shape the growth and physiological traits of their fungal hosts through nutrient transformation, metabolic interactions, synergistic extracellular enzyme activity, and the modulation of environmental adaptation. In edible mushroom cultivation systems, bacterial‒fungal interactions influence hyphal extension, substrate utilization efficiency, and fruiting body formation, underscoring the importance of these bacteria as integral components of mushroom growth and development [8]. Importantly, bacteria detected in fungal cultures are unlikely to be incidental contaminants. Increasing evidence suggests that some bacteria remain tightly associated with fungal hyphae even after isolation and repeated subculturing, notably as endohyphal bacteria or cells stably associated with hyphal surfaces [9]. For example, long-term subcultures of the ectomycorrhizal fungus \u003cem\u003eLaccaria\u003c/em\u003e bicolor have been shown to harbor intracellular \u003cem\u003ePaenibacillus\u003c/em\u003e spp. [10]. In the arbuscular mycorrhizal fungus \u003cem\u003eGigaspora margarita\u003c/em\u003e, the endobacterial symbiont \u003cem\u003eCandidatus Glomeribacter gigasporarum\u003c/em\u003e is vertically transmitted through spores, and its elimination by continuous single-spore propagation markedly affects host spore morphology and hyphal growth [11]. Similarly, in \u003cem\u003ePiriformospora indica\u003c/em\u003e, the endophytic bacterium \u003cem\u003eRhizobium radiobacter\u003c/em\u003e remains stably associated with the host under long-term axenic culture conditions, and a reduction in its abundance markedly impairs host growth, sporulation, and plant growth-promoting capacity [12]. Collectively, these findings suggest that fungus-associated bacteria are not merely incidental occupants but also active contributors to fungal life processes and ecological adaptation. Accordingly, interest in mushroom-associated microbiota has increased steadily in recent years. The bacterial communities associated with cultivated edible mushrooms, including \u003cem\u003eFlammulina filiformis\u003c/em\u003e, \u003cem\u003ePleurotus eryngii\u003c/em\u003e, and \u003cem\u003ePholiota adiposa\u003c/em\u003e, as well as their production environments, have been increasingly investigated, revealing the complexity and dynamism of bacterial assemblages within mushroom cultivation systems [13, 14]. In addition, plate coculture experiments have shown that bacteria and their volatile metabolites can markedly affect the mycelial growth of edible mushrooms [15]. However, previous studies have focused largely on production environments, cultivation substrates, developmental stages, or individual growth-promoting strains, whereas systematic comparisons of hypha-associated bacterial communities across edible mushroom hosts remain scarce. In particular, it remains unclear whether hyphosphere symbiotic bacterial communities are structured by the host taxonomic background and whether such differentiation persists even among closely related hosts. Moreover, the contributions of host physicochemical traits and extracellular enzyme activities to bacterial community assembly and divergence have not been systematically evaluated.\u003c/p\u003e\n\u003cp\u003eHere, we used eight cultivated edible mushroom species as model hosts and performed integrated analyses of hyphal growth traits, physicochemical properties, extracellular enzyme activities, and 16S\u003cem\u003e\u0026nbsp;rRNA\u003c/em\u003e gene amplicon sequencing to systematically characterize the composition and diversity patterns of these mushrooms. We further compared bacterial community variation across six genera and among different strains within species. Spearman correlation analysis together with RDA was then used to assess the contributions of the host taxonomic background and physicochemical traits to bacterial community assembly. This study aimed to elucidate edible mushroom‒bacterium interactions and the ecological principles underlying microbiome assembly, thereby providing a theoretical foundation for efficient mushroom cultivation, targeted microbiome manipulation, and the functional exploitation of mushroom‒associated bacteria\u003c/p\u003e"},{"header":"Materials and Methods","content":"\u003ch3\u003eFungal strains\u003c/h3\u003e\n\u003cp\u003eEight commercially cultivated edible mushroom strains representing eight species were included in this study:\u0026nbsp;\u003cem\u003eFlammulina filiformis\u003c/em\u003e (two strains, including a yellow cultivar),\u0026nbsp;\u003cem\u003eHypsizygus marmoreus\u003c/em\u003e (two strains, including a brown cultivar),\u0026nbsp;\u003cem\u003eLyophyllum decastes\u003c/em\u003e,\u0026nbsp;\u003cem\u003eHymenopellis raphanipes\u003c/em\u003e,\u0026nbsp;\u003cem\u003ePleurotus giganteus\u003c/em\u003e and\u0026nbsp;\u003cem\u003eMacrocybe gigantea\u003c/em\u003e. All the strains were isolated from commercially cultivated fruiting bodies. Pure cultures were established via tissue isolation and maintained on potato dextrose agar (PDA) (Supplementary Fig. 1). Detailed information on the strain origin is provided in Supplementary Table 1.\u003c/p\u003e\n\u003ch3\u003eCulture media\u003c/h3\u003e\n\u003cp\u003eAn enriched potato dextrose agar (PDA) medium was prepared from potato infusions obtained from 200 g fresh potato and supplemented with 20 g glucose, 3 g yeast extract, 3 g peptone, 1.5 g KH\u003csub\u003e2\u003c/sub\u003ePO\u003csub\u003e4\u003c/sub\u003e and 1.5 g MgSO\u003csub\u003e4\u003c/sub\u003e per liter. For the solid medium, 20 g agar was added per liter. The LBL medium was prepared from potato infusions obtained from 200 g fresh potato and supplemented with 20 g lactose, 2 g NH\u003csub\u003e4\u003c/sub\u003eNO\u003csub\u003e3\u003c/sub\u003e, 0.5 g MgSO\u003csub\u003e4\u003c/sub\u003e, 1.5 g KH\u003csub\u003e2\u003c/sub\u003ePO\u003csub\u003e4\u003c/sub\u003e and 0.08 g of bromothymol blue (BTB) per liter; the pH was adjusted to 7.0. The media were autoclaved at 121 °C for 20 min. All the strains were cultured at 25 °C in the dark.\u003c/p\u003e\n\u003ch3\u003eITS amplification and phylogenetic analysis\u003c/h3\u003e\n\u003cp\u003eThe fruiting bodies were cleaned and surface-sterilized, and 5 mm thick tissue blocks were aseptically placed on PDA. The cultures were incubated at 25 °C, and the emerging hyphae were subcultured to obtain pure isolates. Genomic DNA was extracted from fresh mycelia via a commercial kit. The ITS region was amplified via the universal fungal primers ITS4 and ITS5 [16]. After verification by agarose gel electrophoresis, the PCR products were sent for commercial sequencing. Sequences with high-quality chromatograms were aligned via ClustalW [17], and low-quality terminal bases were trimmed. Phylogenetic analysis was performed in MEGA 11.0. A neighbor‒joining tree was constructed via the Kimura two-parameter model, and branch support was assessed with 1,000 bootstrap replicates. The resulting ITS phylogeny was used to infer the taxonomic identities and phylogenetic relationships of the eight strains.\u003c/p\u003e\n\u003ch3\u003eMycelial growth and physicochemical measurements\u003c/h3\u003e\n\u003cp\u003eStrains were cultured on PDA plates at 25 °C in the dark, and the mycelial growth rate (mm day⁻¹) was measured as previously described [18, 19]. Mycelial biomass was determined from liquid cultures. After cultivation, the mycelia were harvested, dried to a constant weight and weighed. Biomass was calculated as follows: Biomass (g 100 ml⁻¹) = mycelium dry weight (g)/culture volume (ml) × 100 [20]. The pH of the fermentation broth was measured via a pH meter.\u0026nbsp;Mycelium total carbon content was determined according to T/NAIA 070-2021 via the potassium dichromate titration method\u0026nbsp;[21]. Mycelium total nitrogen content was determined via the Kjeldahl method with an automatic Kjeldahl nitrogen analyzer according to NY/T 1121.24-2012\u0026nbsp;[22, 23]. Laccase activity was assayed via the ABTS radical cation decolorization method with 2,2′-azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) as the substrate\u0026nbsp;[24]. The crude polysaccharide and soluble protein contents, as well as amylase, cellulase and peroxidase activities, were determined via commercial kits from Enzyme-linked Biotechnology Co., Ltd., according to the manufacturer’s instructions.\u003c/p\u003e\n\u003ch3\u003eDNA extraction and 16S rRNA gene sequencing\u003c/h3\u003e\n\u003cp\u003eTotal DNA was extracted from mycelial samples via a commercial DNA extraction kit [25]. The V3-V4 region of the bacterial 16S \u003cem\u003erRNA\u003c/em\u003e gene was amplified via the universal primer pair 338F (5′-ACTCCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). Each PCR mixture (20\u0026nbsp;μL) contained 4\u0026nbsp;μL of 5\u0026nbsp;×\u0026nbsp;TransStart FastPfu buffer, 2\u0026nbsp;μL of 2.5 mM dNTPs, 0.8 μL of each primer (5 μM), 0.4 μL of TransStart FastPfu DNA polymerase, 10 ng of template DNA, and nuclease-free water to 20 μL\u0026nbsp;[26]. PCR was performed on a GeneAmp 9700 thermocycler (ABI) under the following conditions: 95 °C for 3 min; 27 cycles of 95 °C for 30 s, 55 °C for 30 s and 72 °C for 30 s; and a final extension at 72 °C for 10 min. Amplicons were sequenced on an Illumina MiSeq PE300 platform\u0026nbsp;[27, 28].\u003c/p\u003e\n\u003ch3\u003eSequence processing and community analysis\u003c/h3\u003e\n\u003cp\u003eRaw reads were merged via FLASH [29], and adapter and primer sequences were removed via Cutadapt [30] to obtain high-quality reads. Operational taxonomic units (OTUs) were clustered at 97% sequence similarity, and representative sequences were taxonomically assigned via the SILVA database [31]. Low-abundance OTUs, as well as OTUs assigned to chloroplasts or mitochondria, were removed. The samples were then rarefied to the minimum sequencing depth before downstream analyses. On the basis of the rarefied OTU table, the Chao1, ACE, Shannon, and Simpson indices were calculated to assess alpha diversity. Beta diversity was evaluated via principal coordinate analysis (PCoA), non-metric (NMDS) and permutational multivariate analysis of variance (PERMANOVA) on the basis of Bray‒Curtis distances. The top 10 dominant bacterial phyla and the top 20 dominant bacterial genera were used to characterize the community composition. Differentially abundant taxa among groups were identified via LEfSe (LDA=2, \u003cem\u003ep\u003c/em\u003e\u0026lt;0.05).\u003c/p\u003e\n\u003ch3\u003eStatistical analysis\u003c/h3\u003e\n\u003cp\u003eStatistical analyses were performed using GraphPad Prism. Differences among groups were assessed by one-way analysis of variance (ANOVA), followed by appropriate post hoc tests. Community analyses and visualizations were performed via the Majorbio Cloud Platform (https://www.majorbio.com/). The graphical abstracts were created via Adobe Illustrator 2021.\u003c/p\u003e"},{"header":"Results","content":"\u003ch3\u003eITS-based phylogeny and taxonomic relationships of the fungal strains\u003c/h3\u003e\n\u003cp\u003eFruiting body morphology, together with mycelial growth and colony characteristics on PDA-enriched media, is shown for all the tested strains in Supplementary Fig. 1. ITS-based phylogenetic analysis using reference sequences from GenBank was performed to infer the relationships among the tested strains (Supplementary Fig. 2a). The eight strains were resolved into six phylogenetic clades corresponding to \u003cem\u003eFlammulina\u003c/em\u003e (two strains), \u003cem\u003eHypsizygus\u003c/em\u003e (two strains), \u003cem\u003eHymenopellis\u003c/em\u003e (one strain), \u003cem\u003eLyophyllum\u0026nbsp;\u003c/em\u003e(one strain), \u003cem\u003ePleurotus\u003c/em\u003e (one strain), and \u003cem\u003eMacrocybe\u003c/em\u003e (one strain). This classification was consistent with the observed fruiting body morphology. The yellow and white \u003cem\u003eF. filiformis\u003c/em\u003e isolates, as well as the white and brown \u003cem\u003eH. marmoreus\u003c/em\u003e isolates, represented different strains of the same species.\u003c/p\u003e\n\u003ch3\u003eVariation in host mycelial physicochemical traits\u003c/h3\u003e\n\u003cp\u003eThe nutritional and physiological traits of the eight edible mushroom mycelia are shown in Fig. 1a-i.\u003cem\u003eF. filiformis\u003c/em\u003e_Y presented the highest mycelial growth rate (11.62 mm day⁻¹), followed by \u003cem\u003eF. filiformis\u003c/em\u003e_W (10.01 mm day⁻¹) and \u003cem\u003eH. raphanipes\u0026nbsp;\u003c/em\u003e(9.41 mm day⁻¹) (Supplementary Fig. 2b). Mycelial biomass in liquid culture showed an aimilar pattern, indicating a close association between biomass accumulation and mycelial growth (Supplementary Fig. 2c). Among the tested strains, \u003cem\u003eF. filiformis\u003c/em\u003e_W, \u003cem\u003eH. marmoreus\u003c/em\u003e_W, and \u003cem\u003eH. marmoreus\u003c/em\u003e_B presented relatively high pH values, whereas \u003cem\u003eM. gigantea\u003c/em\u003e presented the lowest pH value (Supplementary Fig. 2d). As shown in Fig. 1a-c, significant differences in total carbon content, total nitrogen content, and the carbon-to-nitrogen ratio were observed among the tested strains. \u003cem\u003eF. filiformis\u003c/em\u003e_Y presented the highest total carbon content, whereas\u003cem\u003e\u0026nbsp;L. decastes\u003c/em\u003e presented the lowest value. The total nitrogen content varied within a relatively narrow range, with \u003cem\u003eM. gigantea\u003c/em\u003e, \u003cem\u003eF. filiformis_W\u003c/em\u003e, and \u003cem\u003eH. marmoreus\u003c/em\u003e_B showing relatively high values, and \u003cem\u003eF. filiformis\u003c/em\u003e_Y and \u003cem\u003eP. giganteus\u003c/em\u003e presented relatively low values. For the carbon-to-nitrogen ratio,\u003cem\u003e\u0026nbsp;F. filiformis\u003c/em\u003e_Y presented the highest value, whereas \u003cem\u003eH. marmoreus\u003c/em\u003e_B, \u003cem\u003eL. decastes\u003c/em\u003e, and \u003cem\u003eM. gigantea\u003c/em\u003e presented relatively low values. The crude polysaccharide content was highest in \u003cem\u003eM. gigantea\u003c/em\u003e (166.65 mg g⁻¹), followed by \u003cem\u003eH. marmoreus\u003c/em\u003e_B (94.72 mg g⁻¹) (Fig. 1d). The soluble protein content ranged from 5.41 to 7.61 mg g⁻¹, with the highest value in \u003cem\u003eH. marmoreus\u003c/em\u003e_B and the lowest in \u003cem\u003eL. decastes\u003c/em\u003e (Fig. 1e). \u003cem\u003eF. filiformis\u003c/em\u003e_Y presented the highest amylase activity, followed by \u003cem\u003eF. filiformis\u003c/em\u003e-W and \u003cem\u003eH. raphanipes\u003c/em\u003e, whereas \u003cem\u003eH. marmoreus\u003c/em\u003e_Y and \u003cem\u003eH. marmoreus\u003c/em\u003e_W presented the lowest activities. (Fig. 1f). The highest cellulase activity was observed in \u003cem\u003eH. marmoreus\u003c/em\u003e_W (11.47 U), whereas the lowest activities were detected in \u003cem\u003eH. marmoreus\u003c/em\u003e_B and \u003cem\u003eM. gigantea\u003c/em\u003e, at 3.20 and 2.40 U, respectively (Fig. 1g). In contrast, a similar trend was observed for laccase and peroxidase activities: both activities were relatively high in \u003cem\u003eL.decastes\u003c/em\u003e,\u003cem\u003e\u0026nbsp;F. filiformis\u003c/em\u003e_Y,\u003cem\u003e\u0026nbsp;F. filiformis\u003c/em\u003e_W, and \u003cem\u003eH. raphanipes\u003c/em\u003e but low in \u003cem\u003eH. marmoreus\u003c/em\u003e_W, \u003cem\u003eH. marmoreus\u003c/em\u003e_B, and \u003cem\u003eM. gigantea\u0026nbsp;\u003c/em\u003e(Fig. 1h-i).\u003c/p\u003e\n\u003ch3\u003eHSBC differed among the six host genera\u003c/h3\u003e\n\u003cp\u003eHigh-throughput sequencing generated 1,523,460 valid reads, which were clustered into 1,028 OTUs. These OTUs were assigned to 25 phyla, 49 classes, 121 orders, 230 families and 527 genera (Supplementary Table 2). Venn analysis revealed that 396 OTUs were shared among the six host genera, whereas \u003cem\u003eFlammulina\u003c/em\u003e, \u003cem\u003eHymenopellis\u003c/em\u003e, \u003cem\u003eHypsizygus\u003c/em\u003e, \u003cem\u003eLyophyllum\u003c/em\u003e,\u003cem\u003e\u0026nbsp;Pleurotus\u003c/em\u003e, and \u003cem\u003eMacrocybe\u003c/em\u003e harbored 2, 4, 23, 7, 2 and 12 unique OTUs (Fig. 3a), respectively. Alpha-diversity analysis revealed that HSBC associated with \u003cem\u003eH. marmoreus\u003c/em\u003e and \u003cem\u003eF. filiformis\u003c/em\u003e_Y presented relatively high richness, whereas \u003cem\u003eH. raphanipes\u003c/em\u003e exhibited the lowest alpha diversity (Fig. 2a-b). The Shannon and Simpson indices suggested broadly similar evenness patterns among the host genera, although \u003cem\u003eHypsizygus\u003c/em\u003e and \u003cem\u003ePleurotus\u003c/em\u003e were more distinct from the other groups (Fig. 2b-c). At the OTU level, PCA and NMDS showed tight clustering within groups, indicating good reproducibility, although a few \u003cem\u003eP. giganteus\u003c/em\u003e and \u003cem\u003eM. gigantea\u003c/em\u003e samples deviated from their group centroids (Fig. 3b-c). At the phylum level, all six host genera were dominated by Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes. Proteobacteria was particularly abundant in \u003cem\u003eL. decastes\u003c/em\u003e (47.15%) and \u003cem\u003eP. giganteus\u003c/em\u003e (44.80%), whereas Firmicutes was most abundant in \u003cem\u003eH. marmoreus\u003c/em\u003e (43.60%) (Fig. 3d). At the genus level, \u003cem\u003eTepidimicrobium\u003c/em\u003e was the dominant genus across all six host genera. \u003cem\u003eGlutamicibacter\u003c/em\u003e was enriched in \u003cem\u003eM. gigantea\u003c/em\u003e, \u003cem\u003eH. marmoreus\u003c/em\u003e and \u003cem\u003eF. filiformis\u003c/em\u003e; \u003cem\u003eAcinetobacter\u003c/em\u003e was abundant in \u003cem\u003eL. decastes\u003c/em\u003e and \u003cem\u003eH. marmoreus\u003c/em\u003e; \u003cem\u003eLactobacillus\u003c/em\u003e was significantly enriched in \u003cem\u003eH. marmoreus\u003c/em\u003e; and \u003cem\u003eSphingomonas\u003c/em\u003e was most abundant in \u003cem\u003eH. raphanipes\u003c/em\u003e (Fig. 3e-f). LEfSe further identified distinct biomarker taxa among host genera, indicating clear host-dependent selection on symbiotic bacterial communities (Fig. 3g).\u003c/p\u003e\n\u003ch3\u003eIntraspecific variation differed between \u003cem\u003eF. filiformis\u003c/em\u003e and \u003cem\u003eHypsizygus marmoreus\u003c/em\u003e\u003c/h3\u003e\n\u003cp\u003eTo further assess intraspecific variation, two \u003cem\u003eF. filiformis\u003c/em\u003e strains and two \u003cem\u003eH. marmoreus\u003c/em\u003e strains were compared. Venn analysis revealed that the two \u003cem\u003eF. filiformis\u003c/em\u003e strains shared 444 genera, with 19 and 35 unique genera in the yellow and white strains, respectively (Fig. 4a). The two \u003cem\u003eH. marmoreus\u003c/em\u003e strains shared 497 genera, with 46 and 19 unique genera among the white and brown strains, respectively (Fig. 4b). No significant differences in alpha diversity were observed between the two \u003cem\u003eF. filiformis\u003c/em\u003e strains (Fig. 4e-h). NMDS further revealed that the hyphosphere bacterial community structure did not differ significantly between the two strains. At the genus and phylum levels, none of the 20 most abundant genera differed significantly between the two strains (Fig. 5a-d). In contrast, the two \u003cem\u003eH. marmoreus\u003c/em\u003e strains differed markedly. Compared with the brown strain, the white strain presented significantly greater alpha diversity indices (Fig. 4i-k). NMDS clearly separated the two groups (Fig. 4c). At the phylum level, no significant differences were observed between the two \u003cem\u003eH. marmoreus\u003c/em\u003e strains (Fig. 5e-f), At the genus level, six taxa, \u003cem\u003eTepidimicrobium\u003c/em\u003e, \u003cem\u003eLactobacillus\u003c/em\u003e, \u003cem\u003eGlutamicibacter\u003c/em\u003e, Ruminococcaceae_UCG-014, \u003cem\u003eRalstonia\u003c/em\u003e and \u003cem\u003eErysipelothrix\u003c/em\u003e, differed significantly between the two strains (Fig. 5g-h).\u003c/p\u003e\n\u003ch3\u003ePhysicochemical traits are associated with bacterial community structure\u003c/h3\u003e\n\u003cp\u003eSpearman correlation analysis revealed distinct associations between host mycelial physicochemical traits and bacterial genera. Hierarchical clustering revealed that TN and TS grouped together, C/N clustered closely with SP, and Amy clustered most closely with Lac, which was further grouped with POD and Cel (Fig. 6).\u0026nbsp;These results suggest that these variables exerted broadly similar effects on genus-level distribution patterns. Several genera, including \u003cem\u003eTepidimicrobium\u003c/em\u003e, \u003cem\u003eFermentimonas\u003c/em\u003e, \u003cem\u003eProteiniphilum\u003c/em\u003e, \u003cem\u003eAmphibacillus\u003c/em\u003e, \u003cem\u003eErysipelothrix\u003c/em\u003e, \u003cem\u003eActinomyces\u003c/em\u003e, PMMR1, \u003cem\u003eRalstonia\u003c/em\u003e, and \u003cem\u003ePelomonas\u003c/em\u003e, were positively correlated with TC, C/N, and SP but negatively correlated with POD, Amy, and Lac. In contrast, \u003cem\u003eBacillus\u003c/em\u003e, \u003cem\u003eBrevundimonas\u003c/em\u003e, and \u003cem\u003eAcinetobacter\u003c/em\u003e were relatively strongly positively correlated with Cel and POD but negatively correlated with C/N. Enterobacter had the strongest positive correlation with SP and was also positively correlated with C/N and Lac but negatively correlated with TN. In addition, \u003cem\u003eGlutamicibacter\u003c/em\u003e was positively correlated with TS, \u003cem\u003eOchrobactrum\u0026nbsp;\u003c/em\u003ewas negatively correlatedwith TN, and \u003cem\u003eMethylobacterium\u003c/em\u003e was negatively correlated with C/N. Overall, genus-specific associations with host physicochemical traits were evident, with \u003cem\u003eEnterobacter\u003c/em\u003e showing the strongest response. RDA revealed that the first two axes explained 21.64% and 11.13% of the total variation, respectively, accounting for 32.77% of the overall variation. TS, TN and POD were associated mainly with the negative side of RDA1, whereas SP, C/N and Lac were associated with the positive side of RDA1; Amy and TC were associated primarily with the negative side of RDA2. \u003cem\u003eM. gigantea\u003c/em\u003e was positioned on the positive side of RDA1, which is consistent with SP, C/N and Lac, whereas \u003cem\u003eH. marmoreus\u003c/em\u003e was distributed in the positive RDA1 and negative RDA2 spaces in association with Lac, Amy and TC. In contrast, \u003cem\u003eF. filiformis\u003c/em\u003e and \u003cem\u003eP. giganteus\u003c/em\u003e were located mainly on the negative side of RDA1, closer to TS, TN and POD. \u003cem\u003eH. raphanipes\u003c/em\u003e clustered near the origin, indicating weaker relationships with the measured physicochemical variables, whereas \u003cem\u003eL. decastes\u003c/em\u003e occupied the negative RDA1 and positive RDA2 space and was separated from most of the environmental vectors\u0026nbsp;(Fig. 7).\u003c/p\u003e"},{"header":"Discussion","content":"\u003ch3\u003eHost taxonomic background drives the differentiation of HSBC\u003c/h3\u003e\n\u003cp\u003eIn the present study, HSBC differed significantly among the host genera, although all the groups were consistently dominated by Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes. This pattern suggests that hyphosphere microbiome assembly is not entirely stochastic but instead reflects the combined effects of core conservation and host specificity within a shared environmental bacterial pool. Similar patterns have also been observed in other studies of fungus-associated microbiomes. Liu et al. [13] used high-throughput 16S \u003cem\u003erRNA\u003c/em\u003e amplicon sequencing to characterize microbial communities associated with \u003cem\u003eF. filiformis\u003c/em\u003e from different production regions and reported a conserved set of dominant taxa, whereas the relative abundances of Pseudomonas and Lactobacillus varied markedly among locations. Similarly, Chen et al. [14]\u0026nbsp;reported that the bacterial community associated with Pleurotus eryngii changed significantly across developmental stages and that some bacterial taxa were positively correlated with the mycelial growth rate, indicating their potential growth-promoting effects. Together, Collectively, these findings indicate that the mushroom-associated microbiota exhibits both stability and ecological plasticity, with bacterial–fungal associations being widespread across phylogenetically diverse fungi rather than exceptional occurrences[32]. Similarly, extraradical hyphae of arbuscular mycorrhizal fungi have been shown to recruit conserved and reproducible bacterial communities, suggesting that fungal hyphae can themselves actively shape the surrounding microbial microenvironment\u0026nbsp;[33]. These observations imply that\u0026nbsp;the\u0026nbsp;host taxonomic background in this context reflects more than differences in identity or nomenclature alone\u0026nbsp;and\u0026nbsp;may represent underlying differences in\u0026nbsp;the\u0026nbsp;host resource supply, metabolic strategy, and ecological niche, all of which can contribute to the differentiation of HSBC.\u003c/p\u003e\n\u003ch3\u003eRelationships between microbiome divergence and host trait variation among closely related hosts\u003c/h3\u003e\n\u003cp\u003eTo further evaluate the effects of host phylogenetic relatedness and host trait variation on the hyphosphere microbiota, we compared the bacterial communities associated with two strains of \u003cem\u003eF. filiformis\u003c/em\u003e and two strains of \u003cem\u003eH. marmoreus\u003c/em\u003e. The results revealed that neither the community structure nor the predicted functions differed significantly between the two \u003cem\u003eF. filiformis\u003c/em\u003e strains. In contrast, the community compositions of the two \u003cem\u003eH. marmoreus\u003c/em\u003e strains differed significantly, and alpha diversity analysis indicated that phylogenetic proximity does not necessarily result in microbiome convergence. Instead, when the host taxonomic background is relatively constant, differences in host phenotype and metabolism may become important drivers of microbiome assembly. This interpretation is consistent with previous studies\u0026nbsp;[34]. reported that bacterial communities associated with different fungal phylogenetic groups differed significantly and that these differences could be partly explained by host chemical traits, particularly pH and the C/N ratio. These results suggest that both the host phylogenetic background and the host physicochemical characteristics jointly shape the assembly of fungus-associated microbiota. Emmett et al. [33] further demonstrated that the bacterial communities associated with the extraradical hyphae of arbuscular mycorrhizal fungi remained highly conserved across different soil backgrounds and nutrient conditions. This finding suggests that stable and long-term interactions may exist between fungal hosts and hypha-associated or hypha-internal microbiota and that coadaptation, or even coevolution, may occur in certain systems. In the present study, the contrasting patterns observed between the \u003cem\u003eFlammulina\u003c/em\u003e and \u003cem\u003eHypsizygus\u003c/em\u003e strains may reflect the hierarchical interplay between phylogenetic constraints and host trait divergence. When key physicochemical traits differ only slightly among closely related hosts, phylogenetic constraints may remain the dominant factor maintaining microbiome conservatism. However, when host physicochemical differences become more pronounced, ecological niche differentiation and the associated environmental filtering may override the effect of phylogenetic similarity, thereby promoting shifts in both community composition and potential functional profiles.\u003c/p\u003e\n\u003ch3\u003eHost regulation of hyphosphere bacterial community assembly\u003c/h3\u003e\n\u003cp\u003eFurther Spearman correlation analysis revealed that host physicochemical traits, including total carbon, crude polysaccharide, soluble protein, pH, and the C/N ratio, were significantly associated with the distribution of dominant bacterial genera and overall community structure. These results suggest that host traits may shape community assembly through both resource supply and environmental filtering. The total carbon, crude polysaccharide, and soluble protein contents were positively correlated with several dominant genera, indicating that increased availability of carbon and nitrogen resources may facilitate the colonization and enrichment of specific bacterial taxa. In contrast, the C/N ratio and the activities of some extracellular enzymes were negatively correlated with those of certain genera, suggesting that shifts in host nutritional conditions and metabolic status may restrict the proliferation of particular bacterial taxa and thereby reshape community composition. Previous hyphosphere studies have shown that carbon-rich compounds released from fungal hyphae can significantly alter the composition and functional potential of surrounding microbial communities, thereby making the hyphosphere a distinct ecological niche relative to the bulk environment [35]. In addition, phosphorus forms, organic phosphorus levels, and hyphal continuity have all been shown to influence the assembly of HSBC [36]. Studies of soil microbial communities have likewise identified pH as a major environmental driver of bacterial community differentiation [37]. Therefore, the coexistence of positive and negative correlations between host physicochemical traits and bacterial taxa suggests that host mycelia may selectively filter hyphosphere bacteria through variations in nutrient inputs, microenvironmental conditions, and substrate transformation. The RDA results provided further support for this interpretation. SP, C/N, and Lac presented similar vector orientations, whereas TC and Amy presented broadly consistent distribution patterns, suggesting that these variables may jointly reflect the resource status and metabolic activity surrounding host hyphae and exert coordinated effects on the bacterial community structure. In contrast, TN, TS, and POD were oriented toward the left side of the ordination space. Notably, the Cel vector was short and close to the origin, indicating a limited contribution to the first two axes and suggesting that cellulose-related processes were unlikely to be the primary drivers of community divergence in this system. This interpretation is consistent with previous studies showing that microbial community composition is closely associated with specific enzyme activities, carbon chemistry, and resource availability and that differences in substrate utilization between bacteria and fungi can further promote niche differentiation and community reorganization [38, 39]. In addition, different hosts appeared to exert varying strengths of selection on hypha-associated bacterial communities. The \u003cem\u003eP. giganteus\u003c/em\u003e group was distributed mainly on the right side of the ordination space and was closely aligned with SP, C/N, TC, Amy, and Lac, suggesting a relatively stable association between community structure and these physicochemical traits. In contrast, the\u003cem\u003e\u0026nbsp;L. decastes\u003c/em\u003e group was distributed mainly in the upper-left region and was clearly separated from \u003cem\u003eP. giganteus\u003c/em\u003e, indicating that different host groups may possess distinct nutrient-use strategies or metabolic characteristics that promote directional differentiation of bacterial communities. Previous studies have shown that bacteria associated with edible mushrooms not only contribute to nutrient supply, mycelial growth, and fruiting body formation but also vary significantly with host type and local environmental conditions [40, 41]. Taken together, these findings suggest that HSBC assembly is jointly driven by the host taxonomic background and host physicochemical traits.\u003c/p\u003e"},{"header":"Conclusion","content":"\u003cp\u003eOur results show that host taxonomic identity underlies the broad differentiation of HSBC across edible mushroom genera, giving rise to microbial assemblages that contain both conserved core taxa and host-specific components. Among closely related hosts, mycelial nutritional and physicochemical traits have emerged as increasingly important drivers of bacterial community reassembly. Together, these findings indicate that hyphosphere microbiome assembly is jointly shaped by host evolutionary history and host-associated environmental factors. This work provides insight into the hierarchical contributions of host taxonomy and host traits to microbiome assembly and offers a theoretical framework for the targeted regulation of beneficial microbiota in edible mushroom cultivation.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eAcknowledgments\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis work was supported by the Project of Modern Agricultural Industrial Technology System of Edible Fungi in Fujian Province.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor Contributions\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll the authors contributed to the study\u0026apos;s conception and design. Mengzhe Gao, Yanfei Xu and Lu Wang; investigation: Mengzhe Gao and Shuting Zhou; writing\u0026mdash;original draft preparation: Shuyu Xing; writing\u0026mdash;review and editing: Sujing Sun: Conceptualization, supervision, resources, funding acquisition, writing\u0026mdash;review and editing. All the authors commented on previous versions of the manuscript and approved its final version.\u003c/p\u003e\n\u003cp\u003eFunding\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eData availability\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eAll the data generated or analyzed during this study are included in this published article and its supplementary information files.\u003c/p\u003e\n\u003cp\u003eConflict of interest\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests.\u003c/p\u003e\n\u003cp\u003eOpen Access\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eValverde ME, Hern\u0026aacute;ndez-P\u0026eacute;rez T, Paredes-L\u0026oacute;pez O (2015) Edible mushrooms: improving human health and promoting quality life. 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Isme j 17:1626-1638. https://doi.org/10.1038/s41396-023-01476-z\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"microbial-ecology","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"meco","sideBox":"Learn more about [Microbial Ecology](https://www.springer.com/journal/248)","snPcode":"248","submissionUrl":"https://submission.nature.com/new-submission/248/3","title":"Microbial Ecology","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false},"keywords":"edible mushrooms, hyphosphere-symbiotic bacteria, host filtering, microbial community assembly, Tricholomataceae","lastPublishedDoi":"10.21203/rs.3.rs-9530892/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-9530892/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"Edible mushroom mycelia harbour diverse symbiotic bacteria that likely contribute to nutrient cycling, yet the factors shaping their diversity and assembly remain poorly understood. We characterized hyphal- symbiotic bacterial communities (HSBC) across eight cultivated Tricholomataceae species using 16S rRNA amplicon sequencing. Alpha diversity varied significantly among hosts, with H. marmoreus and F. filiformis_Y exhibiting the highest richness, whereas H. raphanipes showed the lowest. Proteobacteria, Firmicutes, Actinobacteria, and Bacteroidetes dominated all hosts, while Tepidimicrobium, Glutamicibacter, Acinetobacter, Lactobacillus, and Sphingomonas displayed host-specific patterns. Intraspecific comparisons revealed minimal divergence between two F. filiformis strains, but the two H. marmoreus strains differed markedly in both community composition and diversity. Spearman correlation and redundancy analyses indicated that host mycelial physicochemical traits substantially influenced bacterial community differentiation. Together, these results demonstrate that host taxonomy and mycelial traits jointly shape HSBC assembly and diversity, providing a framework for understanding mushroom–bacterium interactions and promoting beneficial microbiota in cultivation.","manuscriptTitle":"Host Taxonomy and Mycelial Traits Shape Hyphosphere-Symbiotic Bacterial Communities in Cultivated Tricholomataceae Mushrooms","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-05-07 19:00:12","doi":"10.21203/rs.3.rs-9530892/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"reviewerAgreed","content":"129436101624532213064273533281329985994","date":"2026-05-04T15:01:36+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"303459384839484176818653571682559862529","date":"2026-05-04T08:20:45+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"108247043990408539214017323730812273849","date":"2026-05-03T15:07:19+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"163242654053667035425981815908761427299","date":"2026-05-01T08:38:06+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"50261850707361105789008908864465410684","date":"2026-04-29T15:33:54+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2026-04-29T13:52:30+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2026-04-29T06:21:31+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2026-04-29T06:20:52+00:00","index":"","fulltext":""},{"type":"submitted","content":"Microbial Ecology","date":"2026-04-26T09:28:22+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"microbial-ecology","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"meco","sideBox":"Learn more about [Microbial Ecology](https://www.springer.com/journal/248)","snPcode":"248","submissionUrl":"https://submission.nature.com/new-submission/248/3","title":"Microbial Ecology","twitterHandle":"","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"Springer Hybrid","inReviewEnabled":true,"inReviewRevisionsEnabled":false}}],"origin":"","ownerIdentity":"c11f4299-8d98-47d5-b9ad-942c31f11f2d","owner":[],"postedDate":"May 7th, 2026","published":true,"recentEditorialEvents":[{"type":"reviewerAgreed","content":"129436101624532213064273533281329985994","date":"2026-05-04T15:01:36+00:00","index":23,"fulltext":""},{"type":"reviewerAgreed","content":"303459384839484176818653571682559862529","date":"2026-05-04T08:20:45+00:00","index":22,"fulltext":""},{"type":"reviewerAgreed","content":"108247043990408539214017323730812273849","date":"2026-05-03T15:07:19+00:00","index":21,"fulltext":""},{"type":"reviewerAgreed","content":"163242654053667035425981815908761427299","date":"2026-05-01T08:38:06+00:00","index":19,"fulltext":""},{"type":"reviewerAgreed","content":"50261850707361105789008908864465410684","date":"2026-04-29T15:33:54+00:00","index":17,"fulltext":""},{"type":"reviewersInvited","content":"12","date":"2026-04-29T13:52:30+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2026-04-29T06:21:31+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2026-04-29T06:20:52+00:00","index":"","fulltext":""}],"rejectedJournal":[],"revision":"","amendment":"","status":"under-review","subjectAreas":[],"tags":[],"updatedAt":"2026-05-07T19:00:12+00:00","versionOfRecord":[],"versionCreatedAt":"2026-05-07 19:00:12","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-9530892","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-9530892","identity":"rs-9530892","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}