Lysin Motif Effectors Facilitate the Endophytic Colonization of Wheat by Sclerotinia sclerotiorum via Modulating Plant Immunity

ABSTRACT

Sclerotinia sclerotiorum is a necrotrophic fungus that causes severe plant diseases but can also colonize wheat as a symptomless endophyte. However, the mechanism by which its effector proteins regulate this dual lifestyle remains unclear. This study aimed to identify key LysM effectors involved in S. sclerotiorum–plant interactions and elucidate their specific functions. Among seven SsLysM genes, SsLysM6 and SsLysM7 were transcriptionally upregulated during both the pathogenic infection of rapeseed and the endophytic colonization of wheat. The ΔSsLysM6 and ΔSsLysM7 mutants exhibited significantly reduced colonization abilities on both their pathogenic host (rapeseed) and their endophytic host (wheat). Transient expression of SsLysM6 and SsLysM7 in Nicotiana benthamiana leaves demonstrated that both effectors suppress chitin‐induced immune responses, including reactive oxygen species (ROS) bursts, MAPK activation, and immune gene expression. Recombinant SsLysM6 and SsLysM7 proteins exhibited chitin‐binding ability and efficiently suppressed chitin‐triggered ROS bursts in wheat leaves. These findings provide critical insights into the molecular mechanism by which S. sclerotiorum establishes compatible interactions across genetically diverse hosts, highlight the central role of LysM effectors in fungal adaptation to distinct lifestyles, and offer new perspectives on the evolutionary continuum between pathogenic and endophytic fungi.

Two LysM effectors of Sclerotinia sclerotiorum are essential for the endophytic colonization of wheat. SsLysM6/7 inhibit chitin‐induced immune responses and undermine wheat immunity to promote fungal growth.

1. Introduction

Fungal cell walls (FCWs) consist of a matrix of components covalently linked to a fibrous, load‐bearing polysaccharide scaffold that encases the fungal cell (Geoghegan et al. 2017). This structural plasticity plays a critical role in fungal physiology, particularly during host colonization as the FCW represents the primary interface between the pathogen and host (Geoghegan et al. 2017). During plant–fungal interactions, plants rely on pattern recognition receptors (PRRs) to detect fungus‐specific molecular signatures, collectively termed microbe‐associated molecular patterns (MAMPs), including cell wall polysaccharides (e.g., chitin, β‐glucans), membrane sterols (e.g., ergosterol), and secreted proteins (Gong et al. 2020). Recognition of these MAMPs rapidly activates innate immune responses, such as the production of antifungal enzymes (e.g., chitinases, glucanases), antimicrobial secondary metabolites (e.g., phytoalexins), and reactive oxygen species (ROS), providing broad‐spectrum defence (Fujikawa et al. 2012; Oliveira‐Garcia and Deising 2013; Chang et al. 2022; Ngou et al. 2022). To overcome host immune surveillance, phytopathogenic fungi employ stealth strategies to evade immunity by dynamically reconfiguring their cell walls, such as through chitin deacetylation (forming chitosan), reducing β‐glucan content, or synthesizing α‐1,3‐glucan, and secreting effectors to sequester MAMPs directly or suppress host defence (Buscaill and van der Hoorn 2021; Park et al. 2017; Saberi Riseh et al. 2024). Notably, both pathogenic and beneficial fungi can evade host chitin receptor recognition by masking MAMPs with proteinaceous effectors (Buscaill and van der Hoorn 2021).

Chitin perception in plants is mediated by surface‐localized lysin motif (LysM)‐containing receptors, which initiate pattern‐triggered immunity (PTI) upon ligand recognition (Antolin‐Llovera et al. 2014; Sanchez‐Vallet et al. 2015; Rovenich et al. 2016). To counteract this defence mechanism, many fungal pathogens secrete effectors harbouring one or more LysM domains that are structurally analogous to plant chitin receptors. These effectors function by sequestering and masking fungal chitin, thereby preventing its degradation by plant chitinases and suppressing chitin‐induced immune responses through inhibition of chitin recognition or downstream signalling events (Gong et al. 2020; Hacquard et al. 2017; Wawra et al. 2019; Rovenich et al. 2016; de Jonge and Thomma 2009). A well‐characterized example is the LysM effector Ecp6, secreted by the fungal pathogen Cladosporium fulvum, which enhances virulence by disrupting chitin‐triggered host immunity. During infection, Ecp6 scavenges chitin oligosaccharides released from the invading hyphal cell walls, effectively preventing host immune activation (de Jonge et al. 2010). In contrast, Mycosphaerella graminicola produces two LysM effectors, Mg1LysM and Mg3LysM, both of which protect fungal hyphae from plant‐derived hydrolytic enzymes. However, only Mg3LysM suppresses chitin‐induced plant defence responses, highlighting its critical role in PTI evasion through inhibition of chitin signalling (Marshall et al. 2011). Similarly, Colletotrichum higginsianum secretes two extracellular LysM proteins, ChELP1 and ChELP2, during the biotrophic phase of infection. Both proteins exhibit high‐affinity binding to chitin oligomers and polymers in vitro and are required for full pathogenicity.

Notably, these effectors display dual functionality by suppressing chitin‐triggered immune responses while also supporting appressorium development (Takahara et al. 2016). In Botrytis cinerea, the LysM effector BcLysM1 is secreted during the early asymptomatic phase of infection, when it binds chitin and chitin oligosaccharides (COS) to suppress chitin‐triggered immunity and protect mycelia from host chitinases. Additionally, BcLysM1 enhances fungal adhesion to the plant leaf surface (Crumière et al. 2024). Beyond these examples, LysM effectors have been identified in a wide range of foliar pathogens, including the rice blast fungus Magnaporthe oryzae (Slp1), the soil‐borne pathogen Rhizoctonia solani (RsLysM), the vascular wilt fungus Verticillium dahliae (Vd2LysM), and the northern corn leaf blight pathogen Setosphaeria turcica (StLysM) (Dölfors et al. 2019; Kombrink et al. 2017; Mentlak et al. 2012; Gong et al. 2024).

The widespread distribution of LysM effectors among diverse pathogenic fungi suggests that they represent a conserved strategy for suppressing chitin‐triggered immunity across a broad range of hosts. Accumulating evidence further indicates that fungal LysM effectors are essential not only in pathogenicity but also in modulating plant immunity during symbiotic and endophytic interactions. For instance, the arbuscular mycorrhizal fungus Rhizophagus irregularis secretes the LysM effector RiSLM, which binds chitin oligosaccharides to protect fungal cell walls from degradation by plant chitinases, thereby facilitating symbiotic colonization (Schmitz et al. 2019; Zeng et al. 2019). Additionally, the beneficial fungus Trichoderma viride employs the LysM effector Tal6 to sequester N‐acetylglucosamine (GlcNAc) oligomers, interfering with plant immune perception and protecting fungal hyphae from host‐derived chitinases (Romero‐Contreras et al. 2019).

Sclerotinia sclerotiorum is a highly destructive ascomycete fungus with an exceptionally broad host range, infecting more than 700 plant species and causing substantial yield losses in economically important legume and brassicaceous crops, including soybean ( Glycine max ) and oilseed rape ( Brassica napus ) (Liu et al. 2024). Interestingly, recent studies have demonstrated that S. sclerotiorum is also capable of endophytic growth in several monocotyledonous plants, such as wheat ( Triticum aestivum ), rice ( Oryza sativa ), barley ( Hordeum vulgare ), maize ( Zea mays ), and oat ( Avena sativa ). In these hosts, S. sclerotiorum has been reported to confer protection against major diseases, including Fusarium head blight (FHB) and stripe rust. Moreover, this fungus not only promotes wheat growth and yield, but may also provide disease resistance in other cereal crops, such as providing resistance to rice blast under field conditions. Beyond its direct effects on plant growth and immunity, it is plausible that S. sclerotiorum also influences the plant‐associated microbiome, with such microbial interactions potentially contributing to the observed growth promotion and disease resistance (Tian et al. 2023). This dual pathogenic‐endophytic lifestyle probably involves complex molecular crosstalk between the fungus and its host plants (Tian et al. 2020). However, whether LysM effectors participate in S. sclerotiorum–host interactions during either the pathogenic or endophytic phase remains unclear.

To elucidate how LysM effectors modulate chitin‐triggered immune responses during the interaction between S. sclerotiorum and wheat, we systematically identified and characterized members of the LysM effector family in S. sclerotiorum. We demonstrate that SsLysM6 (sscle_06g054140) and SsLysM7 (sscle_07g055920) positively regulate both the endophytic capacity and pathogenicity of S. sclerotiorum by suppressing chitin‐induced immune responses, probably through competitive binding to chitin. Collectively, our findings provide new insights into the functional roles of the SsLysM family, advancing our understanding of the molecular mechanisms underlying S. sclerotiorum pathogenesis and endophytic adaptation.

2. Results

2.1. Genome‐Wide Analysis and Functional Annotation of LysM Domain‐Containing Proteins in S. sclerotiorum

Previous studies have shown that LysM effector proteins are widely distributed among diverse fungal species (Akcapinar et al. 2015; Tian et al. 2022). However, the presence and characteristics of LysM effectors in the S. sclerotiorum genome remain largely uncharacterized. To identify LysM effectors in S. sclerotiorum, a genome‐wide search was performed using the amino acid sequence of Ecp6 as the query. After removing redundant sequences, candidate genes were analysed using SMART for domain validation. Seven proteins containing LysM domains were identified and designated as SsLysM1 to SsLysM7. The predicted SsLysM proteins varied in length, ranging from 228 amino acids (SsLysM7) to 878 amino acids (SsLysM1) (Figure S1A). Among them, SsLysM2 (sscle_08g066840) and SsLysM4 (sscle_06g054180) lack signal peptides (Figure S1A). SsLysM1 (sscle_15g107050), SsLysM3 (sscle_03g024480), SsLysM5 (sscle_12g088330), SsLysM6 (sscle_06g054140), and SsLysM7 (sscle_07g055920) were predicted to be putative effector proteins based on the presence of signal peptides and the absence of transmembrane domains (Figure 1A).

FIGURE 1.

Secreted proteins SsLysM6 and SsLysM7 are upregulated during the colonization of wheat and oilseed rape by Sclerotinia sclerotiorum. (A) Schematic diagrams of SsLysM6 and SsLysM7, highlighting their conserved motifs. (B, C) Relative transcript abundance of SsLysM6 and SsLysM7 during the endophytic colonization of wheat ( Triticum aestivum ) by S. sclerotiorum. (D, E) Relative transcript levels of SsLysM6 and SsLysM7 during the infection of oilseed rape ( Brassica napus ) by S. sclerotiorum. Transcript levels were normalized to the reference gene Sstub. The $2^{-\mathrm{\Delta \Delta }C\mathrm{q}}$ method was used to calculate relative expression, with the 0 h calibrator defined as 1. Points represent three technical reactions from the same cDNA preparation; bars and error bars indicate their mean and technical SD, respectively. Samples from three inoculated leaves were pooled at each time point to generate one composite biological sample. Data are presented descriptively without inferential statistical analysis. (F) The secretory function of the SsLysM6 and SsLysM7 signal peptides was verified using a yeast secretion assay. The yeast strain YTK12 was transformed with either the empty vector pSUC2 (negative control), pSUC2‐SsLysM6SP‐1, pSUC2‐SsLysM6SP‐2, pSUC2‐SsLysM7SP‐1, pSUC2‐SsLysM7SP‐2, or pSUC2‐Avr1SP (positive control). The transformed yeast strains were then tested for growth on CMD − W or YPRAA media and for invertase activity using a colorimetric assay with 2,3,5‐triphenyltetrazolium chloride. The data are shown as means ± SD (n = 3).

2.2. Expression Profiling, Phylogenetic Analysis, and Domain Characterization of SsLysM Family Proteins

The domain architecture of SsLysM proteins was analysed using InterProScan, revealing that effectors in this family harbour variable numbers of LysM domains ranging from one to four. SsLysM2 and SsLysM6 contain four LysM domains, SsLysM3 and SsLysM4 contain three, SsLysM7 contains two, and SsLysM1 and SsLysM5 contain one (Figure 1A and S1A). SignalP predicted no N‐terminal signal peptide in SsLysM2 or SsLysM4, making their secretion through the canonical secretory pathway less probable (Figure S1A). None of the seven SsLysM proteins contained a predicted transmembrane helix (Table S1).

Reverse transcription‐quantitative PCR (RT‐qPCR) profiles obtained from the composite samples showed different expression patterns among the seven SsLysM genes during wheat colonization and oilseed rape infection. SsLysM1 showed a modest increase in relative transcript abundance during wheat colonization and generally reduced abundance during oilseed rape infection (Figure S1B,C). The abundance of SsLysM2 transcripts increased moderately during wheat colonization and showed little variation during oilseed rape infection (Figure S1D,E). Transcript levels of SsLysM3 and SsLysM5 decreased relative to the corresponding 0 h time point under both conditions (Figure S1F,G,J,K). SsLysM4 showed no apparent increase in relative transcript abundance under either condition (Figure S1H,I). By comparison, SsLysM6 and SsLysM7 displayed higher relative transcript abundance than at 0 h at several time points in both host–interaction experiments. During wheat colonization, their transcript levels increased by 7.5‐ and 3.9‐fold at 24 h post‐inoculation, respectively. Expression peaked at 36 h, reaching 8.8‐ and 7.0‐fold that of the corresponding 0 h samples (Figure 1B,C). During oilseed rape infection, SsLysM6 and SsLysM7 expression peaked at 6 h, with increases of 6.2‐ and 2.7‐fold, respectively (Figure 1D,E). The presence of predicted signal peptides, together with the comparatively high relative transcript abundance observed for SsLysM6 and SsLysM7 in both host–interaction experiments, provided a rationale for prioritising these two genes for functional analysis.

To investigate the evolutionary relationships and potential functional divergence of SsLysM proteins, a maximum‐likelihood phylogenetic tree was constructed using the WAG + G + F model, based on amino acid sequences from seven SsLysM proteins of S. sclerotiorum and 26 previously characterized LysM proteins from other fungi (Table S2). These proteins clustered into seven distinct clades (Figure S2). We conducted a comprehensive analysis of the LysM domains in SsLysM6 and SsLysM7, comparing them with the LysM domain of Ecp6. At the primary sequence level, the alignment revealed low amino acid sequence similarity between the two (Figure S3A,B), consistent with the extensive diversification observed within the LysM effector protein family (Gong et al. 2024). However, at the tertiary structure level, the predicted models indicated that these domains adopt the highly conserved typical LysM βααβ fold (Figure S3C). This suggests that the functional core of the LysM domain is primarily determined by its conserved spatial conformation rather than by highly conserved sequences.

2.3. Secretory Functions of the Signal Peptides of SsLysM6 and SsLysM7

To determine whether the signal peptides of SsLysM6 and SsLysM7 possess normal secretory activity, we conducted experimental validation using the yeast secretion trap assay. The YTK12 harbouring pSUC2‐Avr1bsp was used as the positive control, while YTK12 alone and YTK12 carrying the empty pSUC2 vector served as negative controls. YTK12 transformants containing either pSUC2‐SsLysM6sp or pSUC2‐SsLysM7sp were able to hydrolyse disaccharides into glucose, leading to the formation of insoluble red triphenyl formazan from 2,3,5‐triphenyl tetrazolium chloride (TTC). These transformants grew normally on YPRAA medium (Figure 1F). Together, these results demonstrate that the signal peptides of SsLysM6 and SsLysM7 possess secretory activity, suggesting that both proteins can be secreted from S. sclerotiorum cells to perform their biological functions.

2.4. SsLysM6 and SsLysM7 Are Not Essential for Normal Growth and Sclerotia Formation

Previous studies have shown that fungal LysM proteins are not solely involved in modulating plant immunity but can also participate in the regulation of fungal growth and development (Sanchez‐Vallet et al. 2015; Levin et al. 2017). To investigate the potential roles of SsLysM6 and SsLysM7 in fungal growth and development, the two genes were individually disrupted by targeted gene replacement using a hygromycin‐resistance cassette (Figure S4A). Two independent knockout mutants of SsLysM6 (ΔSsLysM6‐29 and ΔSsLysM6‐39) and SsLysM7 (ΔSsLysM7‐5 and ΔSsLysM7‐7) were generated and verified (Figure S4B,C). Complementation strains (comSsLysM6‐39 and comSsLysM7‐14) were generated by introducing the wild‐type (WT) allele of SsLysM6 or SsLysM7 into the ΔSsLysM6‐29 and ΔSsLysM7‐5 via Agrobacterium tumefaciens ‐mediated transformation, respectively (Figure S4D,E). Phenotypic analysis of these strains revealed no detectable differences in vegetative growth, colony morphology, mycelial structure, or sclerotia formation among the wild‐type (1980), ΔSsLysM6‐29, ΔSsLysM6‐39, comSsLysM6‐39, ΔSsLysM7‐5, ΔSsLysM7‐7, and comSsLysM7‐14 strains (Figure 2A). Both the knockout mutants and complementation strains exhibited colony morphology and growth rates comparable to the wild type on potato dextrose agar (PDA) (Figure 2A,D). No significant differences in mycelial morphology were observed between the mutants, complementation strains, and the wild type (Figure 2A). The weight and quantity of sclerotia produced by all mutants and complementation strains were also assessed, revealing that deletion of SsLysM6 or SsLysM7 did not affect sclerotia formation in S. sclerotiorum (Figure 2B,C, Data S1). These results suggest that SsLysM6 and SsLysM7 are dispensable for normal growth and sclerotia formation.

FIGURE 2.

SsLysM6 and SsLysM7 do not contribute to the normal growth and development of Sclerotinia sclerotiorum. (A) Colony morphology, hyphal tip morphology, and sclerotial formation of the wild type (1980, WT), ΔSsLysM6‐29, ΔSsLysM6‐39, comSsLysM6‐39, ΔSsLysM7‐5, ΔSsLysM7‐7, and comSsLysM7‐14 grown on potato dextrose agar (PDA) at 20°C. Colony morphology was photographed at 3 days post‐inoculation (dpi), and sclerotial morphology was recorded at 15 dpi. The scale bar represents 1 cm. (B,C) The weight and number of sclerotia produced by strains in (A). The total sclerotial dry weight per plate (B) and the number of sclerotia per plate (C) were quantified. (D) Growth rate of the WT, ΔSsLysM6‐29, ΔSsLysM6‐39, comSsLysM6‐39, ΔSsLysM7‐5, ΔSsLysM7‐7, and comSsLysM7‐14 strains cultivated on PDA at 20°C. The data are shown as means ± SD (n = 5). ns, no significant difference (p > 0.05).

2.5. SsLysM6 and SsLysM7 Contribute to Endophytic Colonization and Modulation of Wheat Growth

To evaluate the roles of SsLysM6 and SsLysM7 in endophytic colonization by S. sclerotiorum, colonization rates were measured. Endophytic infection was quantified using the chitin‐binding fluorescent marker Alexa Fluor 488 conjugated to wheat germ agglutinin (WGA‐AF 488) as a proxy for fungal biomass. WGA‐AF 488 staining intensities were compared among ΔSsLysM6 and ΔSsLysM7 mutants, the WT strain 1980, and the complementation strains colonizing wheat roots. The result confirmed that ΔSsLysM6 and ΔSsLysM7 mutants colonized wheat roots less efficiently than the wild‐type and complementation strains (Figure 3A,B). To determine whether the reduced WGA‐AF 488 signals in wheat roots would be attributed to strain‐specific differences in staining, we examined and quantified WGA‐AF 488 fluorescence in hyphae of the wild‐type strain 1980, two independent ΔSsLysM6 and ΔSsLysM7 mutants, and their corresponding complementation strains. Under identical culture, staining, and imaging conditions, no significant differences in relative WGA fluorescence intensity were detected among the tested strains (Figure S5A,B). These results indicate that deletion of SsLysM6 or SsLysM7 did not detectably alter WGA‐AF 488 staining of hyphal cell wall chitin under the conditions tested. Therefore, the weaker WGA signals detected in wheat roots inoculated with the knockout mutants are unlikely to result from altered WGA staining efficiency and instead support reduced fungal abundance and impaired endophytic colonization. Additionally, fungal biomass in wheat roots co‐cultured with ΔSsLysM6 and ΔSsLysM7 mutants, the wild‐type, and complementation strains was quantified by qPCR. Consistently, wheat roots co‐cultured with ΔSsLysM6 and ΔSsLysM7 mutants contained less fungal biomass than roots co‐cultured with the wild‐type strain 1980. Importantly, the complementation strains fully restored root colonization (Figure 3C). These results suggest that SsLysM6 and SsLysM7 contribute to promote the endophytic colonization of S. sclerotiorum.

FIGURE 3.

SsLysM6 and SsLysM7 positively regulate the endophytic colonization and growth promotion ability of Sclerotinia sclerotiorum in wheat. (A) Hyphal detection of wild‐type (WT), ΔSsLysM6, ΔSsLysM7 and their complementation strains in wheat roots. Wheat seeds were treated with S. sclerotiorum strains before gemination. Root samples were collected and stained after 7 days post‐germination. Hyphae were visualized as green using Alexa Fluor 488‐wheat germ agglutinin (WGA) staining. The wheat root was visualized as purple using propidium iodide (PI) staining. Scale bars = 200 μm. (B) Fluorescence intensity demonstrating the biomass of S. sclerotiorum strains in wheat roots in (A). Fluorescence intensity was quantified from five root samples using ImageJ. The data are shown as means ± SD (n = 5). (C) Relative intensity of ∆SsLysM6, ∆SsLysM7 mutants, and their complementation strains in wheat following co‐cultivation. Samples were collected after 2 weeks, with three plants pooled per replicate and three replicates per treatment. Relative transcript levels of SsITS during the infection progress of S. sclerotiorum in Triticum aestivum . Reverse transcription‐quantitative PCR was performed and data were normalized to the expression of Tatub. The data are shown as means ± SD (n = 3). (D–G) Plant height, flag leaf length, spike length, and number of seeds per spike of wheat in the WT, ΔSsLysM6, ΔSsLysM7, and complementation strain‐treated group. Statistical significance was assessed by one‐way ANOVA followed by Duncan's multiple range test. Different lowercase letters indicate significant differences (p < 0.05).

To investigate the roles of SsLysM6 and SsLysM7 in modulating the wheat growth promotion by S. sclerotiorum, seeds were treated with ΔSsLysM6 and ΔSsLysM7 mutants, the wild‐type strain, and complementation strains. Plant height, as well as flag leaf length and width, were measured at the early flowering stage. All strains, including ΔSsLysM6 and ΔSsLysM7 mutants, the wild type, and complementation strains, promoted wheat growth. However, the ΔSsLysM6 and ΔSsLysM7 mutants exhibited a reduced growth‐promoting capacity relative to the wild type and complementation strains (Figure 3D, Data S1). At anthesis, the average plant heights of seedlings treated with ΔSsLysM6‐29, ΔSsLysM6‐39, ΔSsLysM7‐5, and ΔSsLysM7‐7 were 45.8 ± 3.5, 42.2 ± 6.7, 42.8 ± 3.1, and 46.4 ± 5.1 cm, respectively. These values were significantly shorter than those of seedlings treated with the wild type, comSsLysM6‐39, and comSsLysM7‐14. Flag leaf lengths of wheat seedlings treated with ΔSsLysM6‐29, ΔSsLysM6‐39, ΔSsLysM7‐5, and ΔSsLysM7‐7 were 11.2 ± 1.6, 10.7 ± 2.1, 12.3 ± 1.9, and 13.8 ± 2.3 cm, respectively, significantly shorter than those of seedlings treated with the wild type (13.9 ± 1.9 cm) (Figure 3E, Data S1). At maturation, spike lengths of seedlings treated with ΔSsLysM6‐29, ΔSsLysM6‐39, ΔSsLysM7‐5, and ΔSsLysM7‐7 were 5.4 ± 0.6, 5.5 ± 0.9, 5.6 ± 0.5, and 5.6 ± 0.5 cm, respectively, significantly shorter than those treated with the wild type, comSsLysM6‐39, and comSsLysM7‐14. The spike length of untreated plants was 5.5 ± 0.6 cm (Figure 3F, Data S1). Consistent with the observations on spike length, the wild‐type strain 1980 and the complementation strains significantly increased the number of grains per spike compared to the untreated control. Grain number per spike did not differ significantly among wheat plants treated with the wild type, deletion, or complementation strains (Figure 3G, Data S1). Thus, no detectable effect of SsLysM6 or SsLysM7 deletion on this yield‐related trait was observed under the tested conditions.

2.6. SsLysM6 and SsLysM7 Contribute to the Biocontrol Activity of S. sclerotiorum

Endophytic growth of S. sclerotiorum can induce endophyte‐mediated resistance in wheat against Fusarium head blight (FHB) (Tian et al. 2020). At the initial bloom stage, wheat spikes from seeds treated with ΔSsLysM6 and ΔSsLysM7 mutants, the wild type, complementation strains, and untreated controls were inoculated with a conidial suspension (10⁶ conidia/mL) of Fusarium graminearum PH‐1 labelled with red fluorescent protein (PH‐1‐RFP). At 4 days post‐inoculation (dpi), FHB symptoms initially appeared on spikelets surrounding the inoculation site and subsequently spread to adjacent spikelets. Seeds treated with ΔSsLysM6 and ΔSsLysM7 mutants, as well as the wild type, conferred significant protection against F. graminearum (Figure 4A,B). However, the expansion of FHB symptoms on spikelets of plants treated with the ΔSsLysM6 and ΔSsLysM7 mutants was more pronounced than on those treated with the wild type (Figure 4A,B).

FIGURE 4.

SsLysM6 and SsLysM7 positively regulate the biocontrol activity of Sclerotinia sclerotiorum in wheat. (A, B) Disease symptoms of Fusarium head blight in wheat panicles after seed treatment with ∆SsLysM6 and ∆SsLysM7 mutants or their complementation strains. Spores of the F. graminearum PH‐1 labelled with RFP was inoculation at the concentration of 10⁶ conidia/mL. The disease symptoms of spikes were photographed after 5 days post‐inoculation (dpi). RFP fluorescence was visualized using a stereoscopic fluorescence microscope. Nine wheat panicles were counted in each treatment. Data are presented as the mean ± SD (n = 9). Scale bars represent 20 mm. (C, D) Disease symptoms of Fusarium crown rot at the stem base of wheat after seed treatment using ∆SsLysM6 and ∆SsLysM7 mutants and their complementation strains. Spores of F. graminearum PH‐1‐RFP were inoculated at a concentration of 10⁶ conidia/mL. Representative images of infected wheat shoots were captured 7 dpi. Lesion areas were quantified using ImageJ. Data are presented as the mean ± SD (n = 22). Scale bars represent 10 mm. Statistical significance was assessed by one‐way ANOVA followed by Duncan's multiple range test. Different lowercase letetrs indicate significant differences (p < 0.05).

Symptom analysis revealed that F. graminearum‐infected plants developed severe Fusarium crown rot (FCR) symptoms (Figure 4C). In contrast, plants treated with ΔSsLysM6 and ΔSsLysM7 mutants, the wild type, or complementation strains, followed by F. graminearum PH‐1 inoculation, exhibited minimal symptoms at 7 dpi (Figure 4C). Disease severity was quantified by measuring FCR lesion lengths on the coleoptiles. Average FCR lesion lengths on coleoptiles treated with ΔSsLysM6‐29, ΔSsLysM6‐39, ΔSsLysM7‐5, and ΔSsLysM7‐7 were 10.4 ± 1.4, 10.9 ± 1.3, 11.0 ± 1.8, and 10.8 ± 1.5 mm, respectively (Figure 4D). These were longer than those measured on coleoptiles treated with the wild type (9.4 ± 1.5 mm), comSsLysM6‐39 (9.3 ± 1.3 mm), and comSsLysM7‐14 (10.0 ± 1.3 mm) (Figure 4D, Data S1). These results suggest that SsLysM6 and SsLysM7 positively regulate the biocontrol activity of S. sclerotiorum towards wheat diseases.

2.7. SsLysM6 and SsLysM7 Suppress Chitin‐Induced Immune Responses in Nicotiana benthamiana

We examined whether SsLysM6 and SsLysM7 are capable of suppressing chitin‐triggered immunity. Confocal microscopy detected SsLysM6‐GFP and SsLysM7‐GFP signals in the extracellular regions following plasmolysis (Figure S6A,B). Transient expression of SsLysM6 and SsLysM7 significantly suppressed the chitin‐induced ROS burst compared to the GFP control in N. benthamiana (Figure 5A). In addition, chitin‐induced MAPK activation was largely suppressed when SsLysM6 and SsLysM7 were transiently expressed (Figure 5B). Immunoblotting with an anti‐GFP antibody detected bands corresponding to SsLysM6‐GFP, SsLysM7‐GFP, and GFP in infiltrated N. benthamiana leaves, confirming the accumulation of these proteins under the assay conditions (Figure 5C). The relative expressions of chitin‐induced immune‐related genes NbERF, NbLOX, NbCYP71D20, NbPR1, NbPR2, and NbPTI5 were repressed by expression of SsLysM6 and SsLysM7 (Figure 5D–I). These results suggest that SsLysM6 and SsLysM7 can suppress chitin‐induced immune responses in N. benthamiana.

FIGURE 5.

SsLsM6 and SsLysM7 suppress chitin‐induced immune responses in Nicotiana benthamiana. (A) SsLysM6 and SsLysM7 inhibit chitin‐induced reactive oxygen species (ROS) burst in N. benthamiana leaves. SsLysM6‐GFP, SsLysM7‐GFP, or an empty vector (EV) was transiently expressed in the leaves of N. benthamiana for 48 h. Relative light units (RLU) were detected, and values represent mean ± SE (n = 12 replicates in each treatment). (B) SsLysM6 and SsLysM7 inhibit chitin‐induced MAPK activation in N. benthamiana. Activated MAPKs were detected with anti‐pERK1/2 antibody. Ponceau S staining of RuBisCO (RBC) served as the loading control. (C) Accumulation of GFP, SsLysM6‐GFP, and SsLysM7‐GFP in infiltrated N. benthamiana leaves. Proteins were detected using anti‐GFP antibody. (D–I) SsLysM6 and SsLysM7 suppress chitin‐induced immune gene expression. N. benthamiana leaves expressing SsLysM6, SsLysM7, or an empty vector (EV) were treated with chitin. Relative expression of immune genes was normalized to NbEF1α, and values represent mean ± SD (n = 3 technical replicates, eight leaf discs as a replicate). At each time point, the GFP‐, SsLysM6‐, and SsLysM7‐expressing groups were compared by one‐way ANOVA followed by Tukey's multiple comparisons test. ns p > 0.05, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

2.8. Chitin‐Binding Specificity and Suppression of Chitin‐Triggered ROS by SsLysM6 and SsLysM7 in Wheat

To examine the polysaccharide‐binding properties of SsLysM6 and SsLysM7, we performed in vitro pull‐down assays using chitin, xylan, and cellulose. Recombinant His‐GST‐SsLysM6 and His‐GST‐SsLysM7 were incubated separately with each insoluble polysaccharide. Following centrifugation, the supernatant and pellet fractions were analysed to detect unbound and substrate‐associated proteins, respectively. The well‐characterized chitin‐binding effector BcLysM1 was included as a positive control, and His‐GST alone was used as a negative control. In the chitin pull‐down assay, BcLysM1, SsLysM6, and SsLysM7 were detected in the pellet fraction, whereas His‐GST was detected only in the supernatant (Figure 6A). These results show that both SsLysM6 and SsLysM7 bind chitin under the conditions tested. By contrast, neither SsLysM6 nor SsLysM7 was detected in the pellet fractions of the xylan or cellulose assays (Figure 6B,C). The presence of BcLysM1, SsLysM6, SsLysM7, and His‐GST in the input samples used for the polysaccharide pull‐down assays was confirmed by SDS‐PAGE and Coomassie Brilliant Blue staining (Figure S6C). Thus, SsLysM6 and SsLysM7 bound selectively to chitin among the polysaccharides examined. Structural modelling suggested different chitin‐binding arrangements for the two proteins. SsLysM7 may form an intermolecular chitin‐binding complex, whereas two of the four LysM domains in SsLysM6 may form an intramolecular binding pocket (Figure S6D).

FIGURE 6.

SsLysM6 and SsLysM7 selectively bind chitin and suppress chitin‐induced reactive oxygne species (ROS) production in wheat. (A–C) Polysaccharide affinity‐precipitation assays of recombinant SsLysM6 and SsLysM7 proteins. Purified His‐GST‐BcLysM1, His‐GST‐SsLysM6, and His‐GST‐SsLysM7 were incubated with chitin beads (A), xylan (B), or cellulose (C). His‐GST and His‐GST‐BcLysM1 were included as the negative and positive controls, respectively. The unbound proteins in the supernatant and the substrate‐associated proteins in the pellet were separated by SDS‐PAGE and visualized by Coomassie Brilliant Blue staining. Red asterisks indicate the expected positions of the corresponding recombinant proteins. (D) Suppression of chitin‐induced ROS production by recombinant SsLysM6 and SsLysM7 in wheat leaves. Chitin was pre‐incubated with 5 μM of purified His‐GST, His‐GST‐SsLysM6, or His‐GST‐SsLysM7 before application to the leaf discs. Treatments without chitin were included to determine background ROS production. Luminescence is shown as relative light units (RLU). Data are presented as mean ± SE from 10 independent leaf discs. RLU values at 5 min, corresponding to the peak of chitin‐induced ROS production, were analysed by one‐way ANOVA followed by Duncan's multiple range test. The experiment was independently repeated three times with similar results.

We next assessed whether SsLysM6 and SsLysM7 suppress chitin‐triggered ROS production in wheat leaves (Figure 6D). In the absence of chitin, the protein buffer, His‐GST, His‐GST‐SsLysM6, and His‐GST‐SsLysM7 treatments produced similarly low background ROS signals. Chitin treatment induced a strong ROS burst in the His‐GST control. Co‐incubation of chitin with either His‐GST‐SsLysM6 or His‐GST‐SsLysM7 significantly reduced ROS production relative to the His‐GST control. These results indicate that both SsLysM6 and SsLysM7 suppress chitin‐triggered ROS production in wheat tissues.

2.9. SsLysM6 and SsLysM7 Are Required for the Full Virulence of S. sclerotiorum

To assess the roles of SsLysM6 and SsLysM7 in pathogenicity, detached and in vivo oilseed rape leaves were inoculated with the wild‐type strain 1980, two independent deletion mutants of each gene, and the corresponding complementation strains. Representative images showed that the deletion mutants produced smaller lesions on detached leaves than the wild‐type and complemented strains (Figure 7A). Statistical analysis revealed that the lesion areas caused by ΔSsLysM6 and ΔSsLysM7 mutants were significantly smaller than those caused by the WT strains (Figure 7C,D). Additionally, comSsLysM6‐39 and comSsLysM7‐14 showed fully restored virulence. Oxalic acid is a critical virulence factor for S. sclerotiorum infectivity. However, we did not find any difference in oxalic acid levels in SsLysM6 and SsLysM7 mutants and complementation strains compared to the wild type, suggesting SsLysM6 and SsLysM7 are not involved in oxalic acid production (Figure 7B). Furthermore, agroinfiltration of N. benthamiana leaves with 35S:SsLysM6 and 35S:SsLysM7 constructs followed by inoculation with S. sclerotiorum resulted in significantly larger lesions compared to that with 35S:GFP (Figure 7E,F). These results suggest that SsLysM6 and SsLysM7 are required for the virulence of S. sclerotiorum.

FIGURE 7.

SsLysM6 and SsLysM7 are key effectors contributing to the pathogenicity of Sclerotinia sclerotiorum to oilseed rape. (A) Disease symptoms of S. sclerotiorum strains inoculated on detached oilseed rape (Brassica napus) leaves. Photos were taken 2 days post‐inoculation (dpi). 1980, wild type. The scale bar represents 1 cm. (B) SsLysM6 and SsLysM7 do not contribute to oxalic acid production. The S. sclerotiorum strains were cultured potato dextrose agar plates supplemented with 0.5% bromophenol blue. Photos were taken after 2 days. The scale bar represents 1 cm. (C) Lesion areas on detached oilseed rape leaves after inoculation were quantified using ImageJ, areas were determined at 2 dpi. At least 16 lesions in each experiment were measured, data are presented as means ± SD. Statistical significance was assessed by one‐way ANOVA followed by Duncan's multiple range test. (D) Lesion areas on in vivo oilseed rape leaves after inoculation were quantified using ImageJ lesion were determined 2 dpi. At least 16 lesions in each experiment were measured, data are presented as means ± SD. Statistical significance was assessed by one‐way ANOVA followed by Duncan's multiple range test. Different lowercase letters indicate significant difference (p < 0.05). (E) Disease symptoms on Nicotiana benthamiana leaves following transient expression of SsLysM6 and SsLysM7 and subsequent inoculation with S. sclerotiorum strain 1980. The scale bar represents 1 cm. (F) Lesion areas on N. benthamiana leaves measured by ImageJ. Data are presented as means ± SD (n = 15). Statistical significance was assessed by one‐way ANOVA followed by Tukey's multiple comparisons test.

3. Discussion

In this study, we identified seven LysM proteins in S. sclerotiorum and found that SsLysM6 and SsLysM7 contribute to both endophytic colonization of wheat and pathogenicity on rapeseed. Both genes were induced during the early stage of wheat colonization and oilseed rape infection. In addition, SsLysM6 and SsLysM7 bound chitin and suppressed chitin‐induced immune responses. These findings support a working model in which S. sclerotiorum secretes SsLysM6 and SsLysM7 during wheat colonization. The two effectors bind chitin fragments and thereby reduce their perception by the wheat chitin receptor complex, probably involving TaCEBiP and TaCERK1 (Figure 8). This local suppression of chitin‐triggered immunity facilitates the establishment of endophytic growth. Stable fungal colonization is subsequently associated with enhanced wheat growth and increased resistance to FHB and FCR. Deletion of either SsLysM6 or SsLysM7 impairs fungal colonization and compromises the growth‐promoting and disease‐protective effects. Thus, SsLysM6 and SsLysM7 are important for establishing a compatible interaction, whereas their direct roles in the subsequent systemic responses remain unclear.

FIGURE 8.

Mechanism of SsLysM6‐ and SsLysM7‐mediated regulation of wheat immunity and endophytic growth of Sclerotinia sclerotiorum. In wheat without treatment with S. sclerotiorum, chitin‐triggered immune signalling associated with pathogenic fungal infection remains at the basal level. Following seed treatment with the wild‐type strain 1980, effectors SsLysM6 and SsLysM7 are secreted and bind chitin fragments at the apoplasts of wheat. This activity is proposed to limit chitin perception by the TaCEBiP–TaCERK1 receptor complex, thereby attenuating chitin‐triggered reactive oxygen species (ROS) production, MAPK activation, and defence‐related gene expression. The resulting local immune suppression provides a favourable environment for endophytic colonization. Established colonization is subsequently associated with enhanced wheat growth and increased resistance to Fusarium head blight (FHB), potentially through systemic defence activation and growth‐related metabolic regulation. In contrast, treatment with either the ΔSsLysM6 or ΔSsLysM7 mutant weakens chitin sequestration, allowing stronger activation of chitin‐triggered immunity and reducing fungal colonization. Nevertheless, both mutants retain partial growth‐promoting and disease‐protective activities, indicating that additional fungal factors contribute to these phenotypes.

Purified SsLysM6 and SsLysM7 bound chitin and suppressed chitin‐induced ROS production in wheat, supporting a role in limiting chitin perception during colonization. This function is consistent with that of LysM effectors from pathogenic fungi, including Ecp6 (de Jonge et al. 2010), Mg3LysM (Marshall et al. 2011), Vd2LysM (Kombrink et al. 2017) and Slp1 (Mentlak et al. 2012), as well as RiSLM (Zeng et al. 2019) from the arbuscular mycorrhizal fungus R. irregularis . Thus, suppression of chitin‐triggered immunity is not restricted to pathogenesis but represents a broader adaptation that enables fungi with different lifestyles to persist in plant tissues. In S. sclerotiorum, SsLysM6 and SsLysM7 may similarly sequester chitin oligomers in the apoplast, reducing their availability to the TaCEBiP–TaCERK1 receptor complex (Liu et al. 2023; Wang et al. 2024) and thereby attenuating early immune responses. Their induction during oilseed rape infection further suggests that these proteins may be deployed during both endophytic and pathogenic growth.

The different domain architectures of SsLysM6 and SsLysM7 may underlie differences in their biochemical and biological activities. SsLysM7 contains two LysM domains. In the three‐domain effector Ecp6, LysM1 and LysM3 undergo chitin‐induced intrachain dimerization to form a deeply buried composite groove with picomolar affinity, whereas the remaining LysM2 domain binds chitin with low‐micromolar affinity (Sánchez‐Vallet et al. 2013). By contrast, structural modelling of four characterized dual‐domain effectors—MoSlp1, Vd2LysM, ChELP1, and ChELP2—did not support the formation of an Ecp6‐like intramolecular composite groove (Mentlak et al. 2012; Takahara et al. 2016; Kombrink et al. 2017; Tian et al. 2022). Biochemical analyses further showed that chitin induces intermolecular association and polymerization of ChELP2, MoSlp1, and Vd2LysM in vitro (Tian et al. 2022). These polymeric complexes have been proposed to precipitate at infection sites and thereby reduce the pool of soluble chitin oligomers available for recognition by plant immune receptors, although their formation in planta remains to be demonstrated (Tian et al. 2022). SsLysM7 may use a related intermolecular mechanism, but its binding affinity, binding stoichiometry, and oligomeric state remain to be experimentally determined. Our structural model places the four LysM domains of SsLysM6 in an arrangement compatible with intramolecular chitin binding; however, modelling alone cannot establish the number, affinity, or stoichiometry of the binding sites. SsLysM6 could therefore contain one composite binding site, multiple nonequivalent sites, or a combination of intra‐ and intermolecular interactions. Whether SsLysM6 and SsLysM7 interact directly, either independently of or in response to chitin, also remains unknown. These possibilities may influence ligand‐length preference, apparent affinity or avidity, binding stoichiometry, and protein retention on chitin‐containing surfaces. Chitin‐induced supramolecular assembly has been shown to retain the single‐domain effector Mg1LysM on fungal cell‐wall chitin and protect hyphae from plant chitinases (Sánchez‐Vallet et al. 2020), while related polymerization has also been observed for Mgx1LysM and several dual‐domain effectors (Tian et al. 2021). Whether SsLysM6 and SsLysM7 similarly form higher‐order assemblies or protect S. sclerotiorum hyphae from plant chitinases remains to be determined.

Purified SsLysM6 and SsLysM7 suppressed chitin‐induced ROS production to comparable extents in both N. benthamiana and wheat leaves. Thus, the more pronounced colonization phenotype associated with ΔSsLysM6 cannot be explained by a detectable difference in ROS suppression under the conditions tested. SsLysM6 was more strongly induced during wheat colonization, suggesting that differences in expression, protein abundance, secretion, stability, or spatial distribution may influence the relative contributions of these effectors in planta. The pull‐down assays were qualitative and used equal protein mass rather than equimolar amounts. Given the different sizes and numbers of LysM domains in SsLysM6 and SsLysM7, these assays do not permit direct comparison of their chitin‐binding affinities or stoichiometry.

The relationship between local immune suppression and enhanced systemic disease resistance by S. sclerotiorum remains to be explored. We propose that SsLysM6 and SsLysM7 initially attenuate chitin perception at the root–fungus interface, preventing an immune response sufficiently strong to eliminate the fungus. Once compatible colonization is established, persistent low‐level fungal signals or colonization‐induced changes in host hormone and defence networks may prime distal tissues against subsequent pathogens. It was shown that S. sclerotiorum can colonize wheat roots and subsequently be detected in stems, flag leaves, and spikes. Endophytic colonization was also associated with altered expression of defence‐ and PTI‐related genes, reduced abscisic acid accumulation, increased jasmonic acid in spikes, and changes in photosynthesis and indole‐3‐acetic acid accumulation (Tian et al. 2020). These findings suggest two non‐mutually exclusive routes linking root colonization to ear protection: systemic movement of the fungus into aerial tissues and transmission of host‐derived mobile signals from roots to shoots. Consequently, the weaker FHB and FCR resistance induced by ΔSsLysM6 and ΔSsLysM7 is most conservatively interpreted as a consequence of reduced endophytic establishment rather than evidence that the two effectors directly activate systemic defence signalling.

Accordingly, our model proposes that SsLysM6 and SsLysM7 bind plant apoplastic chitin oligomers and reduce their perception by the putative TaCEBiP‐TaCERK1 complex, thereby facilitating early endophytic colonization. Stable colonization may subsequently promote wheat growth and defence priming through fungal systemic spread, host‐mediated long‐distance immune signalling, or both. The remaining activity of the deletion mutants may be due to the functional redundancy among LysM family members or compensation by other fungal effectors. Spatial and temporal quantification of fungal biomass in roots, crowns, leaves, and spikes, together with root‐restricted inoculation and analysis of defence hormones and marker genes, will further resolve the proposed root‐to‐ear pathway. Overall, our findings extend the established role of LysM effectors in pathogenic interactions to the beneficial endophytic phase of a schizotrophic fungus and identify SsLysM6 and SsLysM7 as important determinants of the balance between local immune suppression and plant‐wide beneficial effects.

4. Experimental Procedures

4.1. Fungal and Bacterial Strains, Growth Conditions

The wild‐type (WT) S. sclerotiorum strain 1980 was cultured on PDA plates at 20°C and maintained on PDA at 4°C. Knockout mutants of SsLysM6 and SsLysM7 were maintained on PDA plates supplemented with 150 μg/mL hygromycin B, whereas complemented mutants were cultured on PDA containing 250 μg/mL G418. The F. graminearum PH‐1‐RFP strain was cultured on PDA plates at 25°C for approximately 3 days. Mycelial plugs were collected from the actively growing colony margin and transferred to carboxymethyl cellulose (CMC) liquid medium. The cultures were incubated at 25°C with shaking at 190 rpm for 5 days to induce macroconidium production. All expression vectors were introduced into A. tumefaciens GV3101 by electroporation. Bacterial cultures harbouring different vectors were grown in Luria Bertani (LB) medium containing 50 μg/mL rifampicin and appropriate antibiotics at 28°C. N. benthamiana, oilseed rape ( Brassica napus ‘Huashuang 4’), and wheat ( Triticum aestivum ‘Zhengmai 9023’) were grown in a greenhouse at 20°C under a 16 h light/8 h dark photoperiod with 60%–80% relative humidity.

4.2. Multiple Sequence Alignment, Conserved Domain Identification, and Phylogenetic Analysis

The genomic sequences of S. sclerotiorum used in this study were obtained from EnsemblGenomes (https://ensemblgenomes.org/). Accession numbers for all genes and proteins are provided in Table S2. The amino acid sequence of Ecp6 (XP_047764299.1) was used as a query to identify SsLysM effectors in S. sclerotiorum. The TMHMM v. 2.0 (https://services.healthtech.dtu.dk/services/TMHMM‐2.0/) and SignalP v. 6.0 (https://services.healthtech.dtu.dk/services/SignalP‐6.0/) servers were used to predict transmembrane regions and signal peptides. GPI‐SOM (https://mendel.imp.ac.at/gpi/gpi_server.html) was used to predict glycosylphosphatidylinositol (GPI)‐anchored proteins. LysM domains were identified using InterPro (https://www.ebi.ac.uk/interpro/). Multiple sequence alignment was performed using MUSCLE with default parameters on full‐length amino acid sequences of eight SsLysM proteins from S. sclerotiorum and 26 well‐characterized homologues from other fungal species. A maximum‐likelihood (ML) phylogenetic tree was constructed in MEGA 11 using the WAG+G + F model with 1000 bootstrap replicates.

4.3. Functional Evaluation of Signal Peptides

To validate the predicted signal peptides, a yeast invertase secretion assay was performed (Yin et al. 2018). DNA fragments encoding the putative signal peptides were amplified using the primer pairs SsLysM6‐SP‐FP/SsLysM6‐SP‐RP and SsLysM7‐SP‐FP/SsLysM7‐SP‐RP (Table S3). The resulting amplicons were cloned into the pSUC2 vector. Recombinant plasmids containing either the SsLysM6 or SsLysM7 signal peptides were individually transformed into YTK12 yeast cells. A plasmid carrying the Avr1b signal peptide from Phytophthora sojae served as a positive control (Dou et al. 2008). Successful transformation was confirmed by culturing yeast cells on CMD − W medium. YPRAA medium, containing 1% yeast extract, 2% peptone, 2% raffinose, 2 mg/mL antimycin A, and 2% agar, was used to assess the secretory function of the signal peptides. TTC was used to assess sucrase secretion activity in the yeast transformants. Transformants were incubated in 10% sucrose solution at 30°C for 35 min. After incubation, the mixture was centrifuged, and 0.1% TTC reagent was added to the supernatant. The reaction was allowed to proceed at room temperature for 5 min to observe colour changes. A positive reaction was indicated by a colour change from colourless to dark red. The Avr1b signal peptide transformant served as the positive control, and YTK12 yeast cells were used as the negative control.

4.4. RNA Extraction and Gene Expression Analysis

Fungal samples were ground in liquid nitrogen, and total RNA was extracted using TRIzol reagent (NEWBio). Genomic DNA was removed using RNase‐free recombinant DNase I (Vazyme). RNA concentration was measured using a spectrophotometer (DENOVIX). Subsequently, 1 μg of RNA was reverse‐transcribed to synthesize first‐strand cDNA using a reverse transcriptase kit (Vazyme).

To assess the expression profiles of all seven SsLysM genes (SsLysM1–SsLysM7) during the colonization of oilseed rape and wheat, the wild‐type strain was cultured on PDA for 36 h. An aliquot of the axenically cultured mycelia was collected immediately before inoculation and used as the 0‐h calibrator, and the inoculated leaf samples from oilseed rape or wheat plants were collected at 3, 6, 12, 24, 36 and 48 h post‐inoculation. At each time point, samples obtained from three separately inoculated leaves were pooled before RNA extraction to generate one composite biological sample per host at each time point (biological n = 1). Within each host experiment, RT‐qPCR analysis of SsLysM1–SsLysM7 was performed using the same RNA and cDNA preparations derived from the same biological sample set.

RT‐qPCR was performed using 2× ChamQ Blue Universal SYBR qPCR Master Mix (Vazyme), gene‐specific primers (Table S3) and a Bio‐Rad CFX96 Touch Real‐Time PCR Detection System, according to the manufacturer's instructions. Each cDNA sample was analysed in three technical reactions within each RT‐qPCR run. The assays were conducted in two separate RT‐qPCR runs using the same cDNA preparations to assess technical repeatability.

Because the same cDNA preparations were used in both runs, the two runs represented technical repeatability assessments rather than independent biological replication. The first RT‐qPCR run was used to generate the expression profiles presented in the figures, and its complete technical reaction‐level C _q values and normalization calculations are provided in Data S2. The second run was used only as an internal assessment of technical repeatability and was not included in the plotted values or other calculations reported in the manuscript.

Transcript levels were normalized to the S. sclerotiorum β‐tubulin gene (Sstub) using the $2^{-\mathrm{\Delta \Delta }C\mathrm{q}}$ method. For each sample, the arithmetic mean C _q value of the three Sstub technical reactions was calculated as ${\overline{\mathit{Cq}}}_{\mathit{\text{Sstub}},t}=\frac{C{q}_{\mathit{\text{Sstub}},1,t}+C{q}_{\mathit{\text{Sstub}},2,t}+C{q}_{\mathit{\text{Sstub}},3,t}}{3}$. The C _q value of each SsLysM gene was then normalized against the mean C _q value of Sstub to obtain three ΔC _q values from three technical replicates at each time point. For each technical replicate, the ΔΔC _q value was calculated by subtracting the ΔC _q value of the equivalently indexed 0‐h axenically cultured mycelial sample from that obtained at the indicated post‐inoculation time point. Relative expression was then calculated as: $R_{r,t}=2^{-\mathrm{\Delta \Delta }C{q}_{r,t}}$, where $R_{r,t}$ denotes the relative expression value for technical reaction r at time point t. Accordingly, each 0‐h technical reaction yielded a ΔΔC _q value of 0, and all three relative expression values at 0 h were therefore defined as $2^0=1$. The three relative expression values derived from technical replicates are plotted as individual points; their mean and standard deviation (SD) are shown solely to describe technical measurement variation. Consequently, the lack of dispersion among the three 0‐h values arises because these data were normalized to 1 and does not imply a lack of technical variation in the raw C _q measurements. Because only one composite biological sample was available at each time point, the RT‐qPCR data were presented descriptively, and no inferential statistical analyses or post hoc multiple‐comparison tests were performed.

4.5. Agrobacterium‐Mediated Transient Expression and Plasmolysis Assay in N. benthamiana

Binary vectors were introduced into A. tumefaciens GV3101 by electroporation. Individual bacterial transformants were cultured overnight at 28°C in LB medium supplemented with appropriate antibiotics. Cultures were harvested by centrifugation at 4000 rpm for 5 min and resuspended in infiltration buffer containing 10 mM MgCl₂, 10 mM MES, 200 μM acetosyringone. The bacterial suspension was adjusted to an OD₆₀₀ of 1.5 and incubated at room temperature at least 3 h before infiltration. To examine the subcellular localization of SsLysM6 and SsLysM7, their full‐length coding sequences, including the native N‐terminal signal peptides, were fused in frame with GFP at the C‐terminus. The resulting SsLysM6‐GFP and SsLysM7‐GFP constructs were transiently co‐expressed with the plasma membrane marker AtHIR4‐mCherry in N. benthamiana leaves. Agrobacterial suspensions carrying the GFP fusion constructs and AtHIR4‐mCherry were mixed at a 1:1 ratio, with a final OD₆₀₀ of 1.5 for each strain, and infiltrated into the abaxial surface of leaves from 4‐week‐old plants using a needleless syringe. GFP co‐expressed with AtHIR4‐mCherry was included as a control. At 36 h post‐infiltration, fluorescence signals in the abaxial epidermal cells were examined using a confocal laser‐scanning microscope. GFP was excited at 488 nm, and fluorescence was collected at 500–530 nm, whereas mCherry was excited at 561 nm, and fluorescence was collected at 580–620 nm. GFP and mCherry signals were acquired sequentially to minimize spectral overlap.

For plasmolysis assays, infiltrated leaf sections were incubated in 1 M NaCl for 10 min before confocal observation. AtHIR4‐mCherry was used to visualize the position of the plasma membrane before and after plasmolysis. The distribution of SsLysM6‐GFP and SsLysM7‐GFP relative to the retracted AtHIR4‐RFP‐labelled plasma membrane and the cell wall was subsequently examined. All localization assays were independently repeated at least three times using leaves from different plants, and representative images are shown.

4.6. ROS Burst Assay

To assess chitin‐triggered ROS burst in N. benthamiana, the full‐length coding sequences of SsLysM6 and SsLysM7, including their respective native N‐terminal signal peptide sequences, were cloned into the binary vector under the control of the CaMV 35S promoter. The resulting constructs were introduced into A. tumefaciens GV3101 and transiently expressed in N. benthamiana leaves as described above. Leaves infiltrated with A. tumefaciens carrying the GFP construct were used as controls. At 48 h post‐infiltration, 12 leaf discs, each 5 mm in diameter, were collected from comparable regions of the infiltrated leaves for each treatment. Individual leaf discs were placed in separate wells of a 96‐well plate containing sterile distilled water and incubated overnight at room temperature to minimize ROS production caused by tissue excision. The water was subsequently replaced with 100 μL reaction solution containing 5 μM L‐012 (Wako), 10 μg/mL horseradish peroxidase (Sigma‐Aldrich), and 20 μg/mL chitin. Parallel samples receiving the same reaction solution without chitin were included to determine basal ROS production associated with Agrobacterium infiltration and transgene expression. Luminescence was measured immediately at 1 min intervals for 45 min using a GloMax multimode microplate reader. ROS production was expressed as relative light units (RLU). Twelve leaf discs were analysed for each treatment, and the experiment was independently repeated at least three times using leaves from different plants.

Suppression of chitin‐triggered immunity was assessed using an L‐012‐based ROS assay on wheat leaf discs. Briefly, leaf discs were treated with 100 nM chitohexaose ((GlcNAc)₆) pre‐incubated for 6 h with or without 5 μM purified His‐GST, His‐GST‐SsLysM6, or His‐GST‐SsLysM7 proteins. Treatments included chitin alone as a positive control. All experiments were performed with at least 10 biological replicates and repeated three times. Data were analysed using one‐way ANOVA followed by Duncan's multiple range test.

4.7. MAPK Activation Assay

Leaf discs (0.25 cm²) were excised from N. benthamiana leaves at 2 days post‐infiltration and transferred to 12‐well plates containing double‐distilled water for overnight recovery. The leaf discs were then treated with the indicated concentrations of chitin for 0, 5, or 15 min. Total proteins were extracted using protein extraction buffer (20 mM Tris–HCl, pH 7.5, 100 mM NaCl, 1 mM EDTA, 10% glycerol, and 1% Triton X‐100), and samples were boiled at 95°C for 10 min. After centrifugation at 10,000 g for 5 min, the supernatants were collected. Protein samples mixed with SDS loading buffer were separated by 10% SDS‐PAGE. Effector proteins and phosphorylated MAPKs were detected by immunoblotting using anti‐GFP (α‐GFP) and anti‐pERK1/2 antibodies, respectively (Qi et al. 2024; Liu et al. 2024).

4.8. Expression Patterns of N. benthamiana Immune Genes

Plant samples were ground to a powder in liquid nitrogen, and total RNA was extracted using TRIzol reagent according to the manufacturer's instructions, First‐strand cDNA was synthesized from 1 μg of total RNA using a reverse transcription kit (Vazyme). For immune gene expression analysis, total RNA was extracted from N. benthamiana leaves transiently expressing the empty vector (EV) or effector constructs and treated with chitin for 0, 1, or 3 h. The expression levels of immune‐related genes (NbERF1, CYP71D20, NbPti5, NbPR1, NbPR2, NbLOX) were normalized to the reference gene NbEF1‐α. The relative expression of genes was analysed by the relative quantification algorithm of 2^−ΔΔCq. Three technical replicates were set up for each sample.

4.9. Wheat Seed Treatment With S. sclerotiorum and Sowing

To evaluate the effects of wild‐type strains, ΔSsLysM6‐29, ΔSsLysM6‐39, comSsLysM6‐39, ΔSsLysM7‐5, ΔSsLysM7‐7, and comSsLysM7‐14 on wheat growth and disease resistance following endophytic colonization, their endophytic potential was compared. Wheat seeds were washed with tap water, surface‐sterilized with 0.5% NaClO for 10 min, and rinsed three times with sterile water. After sterilization, seeds were soaked in sterile water for 4 h, collected, and air‐dried. Fresh mycelial cultures of the tested strains were harvested from PDA plates and further cultured in 50 mL flasks containing potato dextrose broth on a shaker at 20°C for 4 days to produce hyphal fragment suspensions. Before seed treatment, fungal hyphae were fragmented and resuspended in water, with the suspension adjusted to an OD₆₀₀ of 2.0. The suspension was then mixed with wheat seeds at a rate of 100 mL/kg. After thorough homogenization, the mixture was incubated for 6 h. Inoculated seeds were air‐dried at room temperature for 12 h. As a control, seeds were soaked in sterile water for 6 h and dried using the same procedure. Treated seeds were sown in pots containing a sterile mixture of vermiculite and perlite (3:1) and grown in a greenhouse, with six seedlings per pot. Treated seeds were sown in sterile soil and grown in a greenhouse. After 21 days, PCR analysis was performed to assess endophytic colonization of wheat seedling stems (Tian et al. 2020).

4.10. PCR Detection of S. sclerotiorum

Genomic DNA was extracted from frozen stem tissues or fungal mycelia using the cetyltrimethylammonium bromide (CTAB) method. The primer pair XJJ21/XJJ222 was used to specifically detect S. sclerotiorum (Qin et al. 2010).

4.11. Scelerotinia sclerotiorum Hyphae Staining and Visualization Using Confocal Laser Scanning Microscopy

Wheat root samples were collected at 7 dpi following seeds treatment with the 1980, ΔSsLysM6, ΔSsLysM7, comSsLysM6, and comSsLysM7 strains. The staining procedure was adapted from the protocols of Redkar et al. (2018) and Doehlemann et al. (2011). Briefly, the roots were washed three times with distilled water and incubated in 10% (w/v) potassium hydroxide (KOH) for 10 min to clear the tissues. After KOH treatment, the roots were thoroughly rinsed with phosphate‐buffered saline (PBS, pH 7.4) to neutralize residual alkalinity. The samples were subsequently stained with 10 μg/mL wheat germ agglutinin conjugated to Alexa Fluor 488 (WGA‐AF 488; Thermo Fisher Scientific) and 1 μg/mL propidium iodide (PI; Sigma‐Aldrich) in PBS. Staining was performed for 10 min at room temperature in the dark. Finally, the stained roots were rinsed twice with PBS to remove excess dye and mounted on microscope slides in PBS for immediate confocal observation.

Mycelial plugs of the wild‐type S. sclerotiorum 1980, the ΔSsLysM6‐29 and ΔSsLysM6‐39 mutants, the corresponding complementation strain comSsLysM6‐39, the ΔSsLysM7‐5 and ΔSsLysM7‐7 mutants, and the complementation strain comSsLysM7‐14 were inoculated onto PDA plates overlaid with sterile cellophane membranes. After incubation for approximately 24 h at 20°C, actively growing hyphae were collected from the colony margins. The collected hyphae were incubated with 10 μg/mL WGA‐AF 488. Samples were subjected to vacuum infiltration for 5 min and then incubated at room temperature for 30 min in the dark. The stained hyphae were washed twice with PBS to remove unbound WGA‐AF 488 and subsequently mounted in PBS for microscopy.

Stained root and hyphae samples were examined using a confocal laser scanning microscope (TCS SP8, Leica). The WGA‐AF 488 fluorescence signal, which labels fungal chitin, was excited with a 488 nm argon laser, and emission was collected between 500 nm and 540 nm. The PI fluorescence signal, indicating plant cell walls, was excited with a 561 nm diode‐pumped solid‐state (DPSS) laser, and emission was detected between 580 nm and 630 nm. z‐stack images were captured using a 5× objective lens. For visualization of individual WGA‐AF 488‐stained hyphae cultured in vitro, images were acquired using a 60× objective lens. Maximum intensity projections were generated from the z‐stacks with Application Suite X (LAS X) software (Leica).

4.12. Assessment of Wheat Growth and Yield‐Related Traits Following Treatment With S. sclerotiorum Strains Under Greenhouse Conditions

The effects of SsLysM6 and SsLysM7 on the growth‐promoting activity of S. sclerotiorum in wheat were evaluated following the method described by Tian et al. (2025), with minor modifications. The wild‐type strain 1980, SsLysM6 and SsLysM7 deletion mutants, and their corresponding complementation strains were cultured in 50 mL potato dextrose broth at 20°C with shaking at 190 rpm for 5 days. The mycelia were subsequently collected, washed with sterile distilled water (SDW), and mechanically fragmented. The resulting hyphal fragment suspensions were adjusted to an OD₆₀₀ of 2.0 with SDW. Wheat seeds were surface sterilized in 0.5% NaClO for 10 min, rinsed three times with SDW, and soaked in SDW for 4 h. The seeds were then treated with the hyphal fragment suspensions at a ratio of 1 mL suspension per 5 g seeds and incubated for 6 h at 20°C with static incubation. Seeds treated with an equivalent volume of SDW served as the mock control. Following treatment, the seeds were sown in pots containing a sterilized vermiculite‐perlite mixture 4:1 v/v. The plants were maintained in a greenhouse at 20°C, 16 h light/8 h dark photoperiod. Pots were randomly arranged and periodically repositioned within the greenhouse to minimize positional effects.

Growth‐related traits were measured individually at the anthesis stage. Plant height was measured from the soil surface to the tip of the main spike, excluding the awns. Flag leaf length was measured from the ligule to the leaf tip. Spike length was measured from the base to the tip of the main spike, excluding the awns. At physiological maturity, the main spike from each plant was harvested separately, and the number of fully developed seeds per spike was determined by manual counting. Measurements from individual plants were retained as independent raw data points. Data are presented as the mean ± SD. Statistical differences among treatments were evaluated using one‐way ANOVA followed by Duncan's multiple range test.

4.13. Pathogenicity Assays

To evaluate virulence, 0.5‐mm‐diameter mycelial plugs were excised from the colony margins of 48‐h‐old PDA cultures of the wild‐type strain 1980, ΔSsLysM6‐29, ΔSsLysM6‐39, comSsLysM6‐39, ΔSsLysM7‐5, ΔSsLysM7‐7, and comSsLysM7‐14. The plugs were inoculated onto detached leaves of B. napus or on leaves attached to intact plants. Lesion areas were recorded at 48 h post‐inoculation. The detached‐ and intact‐leaf assays were each performed three times.

4.14. Gene Substitution and Complementation in S. sclerotiorum

Knockout mutants of the SsLysM6 and SsLysM7 genes in S. sclerotiorum were generated using the split‐marker recombination method (Catlett et al. 2003). The knockout strategy is illustrated in Figure S4A. Two fragments, each about 1 kb in length and designated as SsLysM6/SsLysM7‐5′ and SsLysM6/SsLysM7‐3′, were amplified from genomic DNA by PCR using primers F1/R1 (containing SalI sites) and F3/R3 (containing XbaI sites) with Fast Pfu polymerase (TransStart) (Figure S4). The PCR products were then inserted into the SalI and XbaI sites of the pUCH18 vector containing a hygromycin resistance cassette. After successful cloning, the constructs pUCH18‐SsLysM6/SsLysM7‐5′ and pUCH18‐SsLysM6/SsLysM7‐3′ were obtained. The fusion fragments SsLysM6/SsLysM7‐5′ and SsLysM6/SsLysM7‐3′, together with two truncated hygromycin resistance genes (SsLysM6/SsLysM7‐5′‐HY and YG‐SsLysM6/SsLysM7‐3′), were amplified using primers F1/HY and YG/R3, respectively (Figure S4). Purified DNA fragments (SsLysM6/SsLysM7‐5′‐HY and YG‐SsLysM6/SsLysM7‐3′; ≥ 10 μg each) were mixed in equimolar amounts and used for transformation to generate SsLysM6/SsLysM7 knockout mutants.

To generate complementation strains, the full‐length SsLysM6 or SsLysM7 gene, including its native promoter region, was amplified from the genomic DNA of wild‐type strains, producing the fragments 5′‐XhoI‐promoter‐SsLysM6‐SacI‐3′ and 5′‐XhoI‐promoter‐SsLysM7‐SacI‐3′. Each fragment was inserted into the pCETNS vector containing a geneticin resistance expression cassette. The promoter‐SsLysM6/SsLysM7 fragment linked to the geneticin resistance cassette was amplified from the vector using Fast Pfu DNA polymerase (TransStart), purified, and used (≥ 10 μg) for transformation of protoplasts derived from the SsLysM6/SsLysM7 knockout strain. Transformation of S. sclerotiorum protoplasts was performed using the polyethylene glycol‐mediated method as described by Liu et al. (2024). All primers used for mutant verification and genetic complementation assays are listed in Table S3.

4.15. Protein Purification

The coding sequence for the LysM effector protein (BcLysM1, SsLysM6 and SsLysM7) was amplified and cloned into the pGEX4T‐1 vector for expression in Escherichia coli DE3 cells. Protein expression was induced with 0.1 mM isopropyl β‐D‐1‐thiogalactopyranoside (IPTG) at an optical density (OD₆₀₀) of 0.6–0.8, followed by incubation at 16°C overnight. Cells were harvested by centrifugation and lysed via sonication in a buffer containing 50 mM Tris–HCl (pH 7.5), 300 mM NaCl, and 5 mM imidazole. The recombinant His‐GST‐tagged protein was purified with Ni‐NTA sepharose 6FF (BBI Solutions). The eluted protein was dialysed against PBS and concentrated using an Amicon Ultra centrifugal device (Millipore). Protein concentration was determined using the DeNoVIX assay, with bovine serum albumin (BSA) as the standard.

4.16. Chitin Binding Affinity Precipitation Assays

The polysaccharide‐binding activities of SsLysM6 and SsLysM7 were examined using an affinity precipitation assay adapted from Gong et al. (2024). Purified recombinant proteins His‐GST‐SsLysM6, His‐GST‐SsLysM7, His‐GST‐BcLysM1, and His‐GST were adjusted to 30 μg/mL in 500 μL of chitin‐binding buffer containing 20 mM Tris–HCl, pH 8.0, 500 mM NaCl, 1 mM EDTA, and 0.05% Tween 20. Each protein was incubated separately with 500 μL of chitin magnetic bead suspension (New England Biolabs), 3 mg xylan (Sigma), or 3 mg cellulose (Aladdin). The mixtures were incubated at 4°C for 6 h with continuous gentle agitation. Following incubation, the insoluble matrices were collected by centrifugation at 13,000 g for 5 min. The supernatants containing unbound proteins were carefully collected. The resulting pellets were resuspended in 100 μL of chitin‐binding buffer. The pellet and supernatant fractions were then mixed with SDS‐PAGE sample buffer and heated at 100°C for 10 min. Proteins in the bound pellet and unbound supernatant fractions were analysed by SDS‐PAGE followed by Coomassie Brilliant Blue staining. The experiments were independently repeated three times.

4.17. FHB Infection Assay and Quantification of PH‐1‐RFP Colonization

FHB infection assays were performed using a single‐floret inoculation method as described previously (Hao et al. 2023), with minor modifications. Macroconidia of the RFP‐expressing F. graminearum PH‐1‐RFP were produced in CMC liquid medium. At the heading stage, 10 μL of a conidial suspension containing 10⁶ conidia/mL was introduced between the lemma and palea of a single floret located in the fourth or fifth spikelet from the base of each wheat spike. Following inoculation, the spikes were covered with transparent plastic bags to maintain high relative humidity and incubated at 25°C. The bags were removed at 36 h post‐inoculation. At least 10 spikes from independent plants were inoculated for each seed treatment group. At 7 dpi, PH‐1‐RFP colonization was evaluated by fluorescence imaging. All spikes were imaged using identical acquisition parameters. RFP fluorescence was quantified using ImageJ; RFP fluorescence intensity was used as a relative proxy for PH‐1‐RFP colonization in wheat spikes. Measurements obtained from individual wheat spikes were treated as independent biological observations and used for statistical comparisons among seed treatment groups.

4.18. Wheat Coleoptile Infection Assay With F. graminearum Following Seed Treatment With S. sclerotiorum

Wheat seed surface sterilization and treatment with hyphal fragment suspensions of the indicated S. sclerotiorum strains were performed as described above. Seeds treated with SDW served as the mock control. The treated seeds were transferred to sterile germination boxes containing sterile filter paper moistened with SDW. The boxes were maintained at 20°C under a 16 h light/8 h dark photoperiod. At 7 days after germination, the upper 2–3 mm of each coleoptile was removed with sterile scissors. A 10 μL aliquot of an F. graminearum PH‐1‐RFP conidial suspension at 10⁶ conidia/mL was applied to the cut surface. The seedlings were maintained under 25°C. Coleoptile lesions were photographed at 7 dpi. Lesion areas were quantified using ImageJ and the lesion area was calculated after spatial calibration. Each seedling was treated as an independent biological replicate.

4.19. Quantification and Statistical Analysis

Data for quantification analyses are presented as mean ± SD or mean ± standard error SE. The number of repeats is shown in the figure legends. The statistical analyses were performed by Student's t‐test or one‐way ANOVA in GraphPad Prism v. 8.0 or SPSS. Quantification of immunoblotting signals was performed using ImageJ software.

Author Contributions

Yamin Du: investigation, writing – original draft, data curation, validation, methodology, software, visualization. Haohao Ren: investigation. Hui Tang: investigation. Yafei Chen: funding acquisition, investigation. Jianqi Wang: investigation. Xiaofan Liu: investigation. Jiatao Xie: formal analysis, methodology. Jiasen Cheng: formal analysis, methodology. Yanping Fu: formal analysis. Daohong Jiang: formal analysis. Bo Li: conceptualization, formal analysis. Xiao Yu: supervision, writing – review and editing, project administration, conceptualization, funding acquisition, resources.

Funding

This research was supported by the National Natural Science Foundation of China (grant nos 32302463 to Y.C. and 32572937 to X.Y.), Fundamental and Interdisciplinary Disciplines Breakthrough Plan of the Ministry of Education of China (JYB2025XDXM703), and Fundamental Research Funds for the Central Universities (2662025ZKPY008).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Data S1: Raw numerical data used for the statistical analyses related to Figures 2C, 3D–G and 4D.

Data S2: Raw C _q values and calculated relative‐expression values used for reverse transcription‐quantitative PCR analysis related to Figure 1 and Figure S1.

Figure S1: Domain organization and host‐induced expression patterns of five SsLysMs during wheat ( Triticum aestivum , Ta) and oilseed rape ( Brassica napus , Bn) colonization. (A) Schematic diagram illustrating the protein domain architectures of five SsLysMs (SsLysM1 to SsLysM5). Grey bars denote N‐terminal signal peptides, green rounded rectangles represent conserved LysM carbohydrate‐binding domains, and the total amino acid length of each full‐length protein is labelled on the right‐hand side of each schematic. (B, D, F, H, and J) Relative transcript levels of SsLysM1‐SsLysM5 during the endophytic colonization of wheat by Sclerotinia sclerotiorum. (C, E, G, I, and K) Relative transcript levels of SsLysM1–SsLysM5 during infection of oilseed rape by S. sclerotiorum. For each time point, samples obtained from three separately inoculated leaves were pooled to generate one composite biological sample. Axenically cultured S. sclerotiorum mycelia collected before exposure to hosts were used as the 0‐h calibrator. Transcript levels were normalized against Sstub and calculated using the 2^−ΔΔCq method, with all three technical‐reaction‐level values at 0 h set to 1. Individual points represent three technical reactions performed using the same cDNA preparation; bars and error bars indicate their mean and technical SD, respectively. The data are presented descriptively without inferential statistical analysis.

Figure S2: Proteins containing LysM domains are widely distributed across diverse fungal species. Phylogenetic analysis of the proteins containing LysM domains of Scelrotinia sclerotiorum and their homologues in various fungal species. The phylogenetic tree was constructed using MEGA 11 with the WAG + G + F model. Bootstrap support values (1000 replicates) greater than 70% are shown at each node.

Figure S3: Sequence alignment, conserved motifs, and structural characterization of SsLysM6, SsLysM7 and reference Ecp6 protein. (A) Multiple sequence alignment of individual LysM motifs from Ecp6, SsLysM6 and SsLysM7. Conserved amino acid residues are shaded in light blue. Residue positions are labelled above the alignment. (B) Sequence logo showing the frequency and conservation of amino acids across aligned LysM sequences. The height of each character corresponds to the conservation score at that position. (B) Sequence logos of two relatively conserved regions identified from the multiple sequence alignment of the full‐length amino acid sequences of Ecp6, SsLysM6 and SsLysM7. The upper and lower logos represent conserved regions I and II, respectively. At each position, the total height of the amino acid stack indicates sequence conservation, whereas the height of each letter reflects the relative frequency of the corresponding amino acid. (C) Representations of the predicted three‐dimensional structures of Ecp6, SsLysM6 and SsLysM7. Homology models of the three proteins were generated using SWISS‐MODEL. Ecp6 contains three LysM domains, SsLysM6 contains four LysM domains, and SsLysM7 contains two LysM domains. Individual LysM domains are labelled and coloured separately: LysM1, pink; LysM2, green; LysM3, light blue; and LysM4, beige. The predicted structures were visualized and rendered as cartoon representations using PyMOL.

Figure S4: Gene deletion mutants of SsLysM6 and SsLysM7 were confirmed by PCR analysis. (A) Gene knock‐out strategy for the SsLysM6 and SsLysM7. The gene coding region was replaced with the hygromycin phosphotransferase gene cassette (HYG) by homologous recombination. (B, C) The deletion mutants were verified by PCR amplification using specific primer pairs. The full‐length HptII fragment was amplified using primers F2 and R2. The genomic region upstream of the PtrpC promoter overlapping SsLysM6/SsLysM7 was amplified using primers F1 and HY. While the trpC terminator overlapping the downstream region of SsLysM6/SsLysM7 was amplified using primers YG and R3. The full‐length sequences of SsLysM6 and SsLysM7 were amplified using primer pairs SsLysM6‐FP/SsLysM6‐RP and SsLysM7‐FP/SsLysM7‐RP, respectively. (D, E) Reverse transcription PCR to detect the mRNA levels of SsLysM6 or SsLysM7 in ΔSsLysM6, ΔSsLysM7, and the respective complemention strains.

Figure S5: Knockout of SsLysM6 or SsLysM7 does not alter the WGA‐binding capacity of Sclerotinia sclerotiorum vegetative hyphae. (A) Representative laser confocal micrographs of S. sclerotiorum hyphae grown on potato dextrose agar plates stained with WGA‐Alexa Fluor 488 (green signal). The tested strains consist of wild‐type 1980, two independent SsLysM6 knockout mutants (ΔSsLysM6‐29, ΔSsLysM6‐39), its corresponding complemented strain comSsLysM6‐29, two independent SsLysM7 deletion mutants (ΔSsLysM7‐5, ΔSsLysM7‐7) and the complemented strain comSsLysM7‐14. Scale bar = 20 μm for all images. (B) Quantitative analysis of relative WGA fluorescence intensity among the indicated S. sclerotiorum strains. Data are presented as mean ± SD (n = 16). Statistical significance was assessed by one‐way ANOVA followed by Duncan's multiple range test.

Figure S6: Subcellular localization of SsLysM6 and SsLysM7 in Nicotiana benthamiana epidermal cells and structural modelling of chitin‐binding domains of LysM proteins. (A) Confocal laser scanning microscopy of N. benthamiana epidermal cells transiently co‐expressing GFP, SsLysM6‐GFP, or SsLysM7‐GFP with the plasma membrane marker AtHIR4‐mCherry before plasmolysis. The full‐length SsLysM6 and SsLysM7 proteins contained their native N‐terminal signal peptides, and GFP was fused to their C‐termini. GFP alone served as the control. (B) Subcellular distribution of the indicated proteins following plasmolysis induced by treatment with 1 M NaCl for 10 min. White arrowheads indicate GFP fluorescence retained between the retracted mCherry‐labelled plasma membrane and the cell wall. GFP, green fluorescent protein channel; mCherry, plasma membrane marker channel; BF, bright‐field image; Merge, merged fluorescence and bright‐field images. Scale bars = 36.8 μm. (C) SDS‐PAGE analysis of the recombinant proteins used as inputs for the in vitro polysaccharide pull‐down assays. Purified His‐GST‐BcLysM1, His‐GST‐SsLysM6, His‐GST‐SsLysM7, and His‐GST were separated by SDS‐PAGE and visualized by Coomassie Brilliant Blue staining. Red asterisks indicate the corresponding recombinant protein bands. Molecular mass markers are indicated on the left. (D) Structural comparison of Ecp6 with predicted models of SsLysM6 and SsLysM7. The individual LysM domains are shown as surface representations: LysM1, pink; LysM2, green; LysM3, light blue; and LysM4, beige. Chitin is shown as cyan sticks. The Ecp6 structure illustrates the established composite chitin‐binding groove formed by LysM1 and LysM3, whereas the SsLysM6 and SsLysM7 complexes represent computationally predicted docking poses. Chitin docking for Ecp6, SsLysM6 and SsLysM7 was performed using cb‐dock2, and the resulting complexes were visualized using PyMOL.

Table S1: Predicted signal peptides and transmembrane helices in the seven SsLysM proteins.

Table S2: The information of fungal LysM proteins related to SsLysMs.

Table S3: List of primers used in the study.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.