Domain gain or loss in a fungal chitinase enables specialization towards antagonism or immune suppression

Domain: neuroscience
Published: 2026-03-30
Editorial outcome: major_revision

Abstract
Effector proteins orchestrate interactions that determine host compatibility and microbial competition. Among these are microbial chitinases, widespread enzymes involved in nutrient acquisition, fungal cell-wall remodelling, and antagonism, yet how their functions diversify across ecological roles remains unclear. Here, we show that modular domain variation within a conserved GH18 chitinase family enables specialization between microbial antagonism and host immune suppression in the beneficial root endophyte Serendipita indica. The chitinase SiCHIT, bearing a C-terminal CBM5 domain, is induced during fungal confrontation and inhibits growth of the phytopathogen Bipolaris sorokiniana, thereby protecting host roots. Deleting CBM5 abolishes this antagonistic activity, whereas grafting CBM5 onto its paralog SiCHIT2, which lacks this domain, confers antifungal function. In contrast, SiCHIT2 is induced during root colonization and suppresses chitin-triggered immunity, promoting host compatibility. These results show how domain loss and regulatory divergence can reprogram antimicrobial enzymes into immune-suppressive effectors in beneficial plant-associated fungi.

Introduction
Microbial effector proteins shape the outcome of host–microbe and microbe–microbe interactions across diverse biological systems. A central question in molecular ecology and symbiosis research is how new ecological functions emerge from existing protein architectures1. Structural modularity enables proteins to acquire new functions by altering domain composition through duplication, gain and loss, or recombination2,3. In microbial effectors, such architectural variations potentially drive shifts in ecological roles, including inter-microbial antagonism or host immune suppression. However, direct mechanistic evidence linking domain changes to functional repurposing remains limited, particularly in mutualistic fungi.

Mutualistic fungi are ecologically important members of the rhizosphere microbiome, where they support plant health and shape microbial community structure and function4. This diverse group includes ectomycorrhizal and arbuscular mycorrhizal fungi, as well as beneficial root endophytes. Unlike mycorrhizal fungi, root endophytes often colonize plant roots without forming specialized structures and, depending on the host and environmental context, can enhance plant performance by promoting nutrient uptake, water acquisition, tolerance to abiotic stress, and suppression of microbial pathogens that threaten the root niche5–10.

These fungi exemplify the evolutionary plasticity of fungal lifestyles, flexibly occupying ecological niches along the saprotrophy-to-symbiosis continuum. To colonize plant roots and maintain their niche, beneficial root endophytes must overcome two key challenges: evasion of host immunity and competition with microbial antagonists and plant pathogens11,12. This dual challenge is increasingly understood as part of a broader ecological strategy, supported in some cases by synergistic interactions with bacterial members of the core microbiota8,13. Both immune evasion and microbial competition are mediated by secreted effector proteins11,14–17. Effector-mediated immune evasion often involves the degradation or masking of microbe-associated molecular patterns (MAMPs), such as β-1,3/1,6-glucans and chitin, to prevent recognition by host pattern recognition receptors18–23. In parallel, effectors can modulate the surrounding microbiota by suppressing microbial competitors, a strategy well documented in plant pathogens11,14,16 and potentially present in mutualistic fungi as a mechanism for niche defense.

The beneficial root endophyte Serendipita indica (Si), a member of the widespread fungal order Sebacinales (Basidiomycota), colonizes a broad range of host plants and is known for its ability to promote growth and suppress plant immunity while protecting the host from pathogens8,24–29. Colonization is confined to the rhizoplane, epidermis, and cortex, and depends on an intact host immune system30. In addition to its host-associated functions, Si exhibits strong antifungal activity in the rhizosphere, particularly against the pathogen Bipolaris sorokiniana (Bs), the causal agent of spot blotch and root rot in cereals8,25,31. This antagonism involves multiple molecular factors, with a GH18-CBM5 endochitinase (SiCHIT) playing a prominent role by restricting Bs growth and recapitulating the protective effect of Si in planta31.

GH18 chitinases, found across all fungal phyla, cleave β-1,4-glycosidic bonds in chitin, a major structural component of fungal cell walls. These enzymes fulfill diverse roles in nutrient scavenging, hyphal remodeling, and fungal antagonism32,33. Their catalytic activity depends on a conserved DxxDxDxE motif, which defines the substrate-assisted catalytic mechanism characteristic of this family34,35. While some chitinase-derived effectors have lost enzymatic activity and evolved to sequester MAMPs36, others retain catalytic function and are functionally enhanced by accessory domains such as carbohydrate-binding modules (CBMs)37–39. These non-catalytic domains increase substrate affinity and specificity, particularly for crystalline or polymeric forms of chitin40,41. SiCHIT features a C-terminal CBM5 domain, a modular architecture found predominantly in bacteria and Basidiomycota but absent in Ascomycota, suggesting a lineage-specific evolutionary adaptation31,33.

Here, we investigate the functional significance of CBM5 in SiCHIT and examine how domain architecture modulates effector activity, enabling a switch between microbial antagonism and immune evasion. We show that the CBM5 domain is essential for the antifungal and plant-protective function of SiCHIT. This enzyme is specifically expressed during fungal competition and restricts the growth of the phytopathogen Bs in the rhizosphere, thereby protecting host roots. In contrast, its closely related paralog SiCHIT2 lacks CBM5 and is induced during root colonization, where it suppresses chitin-triggered immune responses by preventing host-derived reactive oxygen species accumulation. Through domain-swapping experiments, we demonstrate that CBM5 functions as a modular determinant capable of switching a chitinase’s ecological role from microbial antagonism to host immune suppression, while retaining its core chitinolytic activity. Together, our findings support a model in which effector specialization in mutualistic fungi may arise through gene duplication, domain loss, and transcriptional divergence, enabling the functional repurposing of an antimicrobial enzyme for host adaptation.

Results
CBM5 enhances binding to crystalline chitin and fungal cell wall without altering product specificity
To investigate the role of the C-terminal CBM5 domain in SiCHIT (PIIN_03543; Pirin1_74346), we generated three recombinant variants, all lacking their native signal peptides: full-length SiCHIT, a catalytically inactive mutant (SiCHITE196Q), and a truncated version lacking the CBM5 domain (SiCHIT-CBM5), which retained the proline-rich linker. The truncated construct was produced by excluding the final 141bp from the coding sequence. All recombinant proteins were purified via affinity chromatography and verified by SDS–PAGE (Supplementary Fig. 1).

We first assessed chitinase activity on insoluble crab shell chitin by quantifying reducing sugars after 20h of incubation (Fig. 1A). This assay revealed no major differences between full-length SiCHIT and SiCHIT−CBM5. However, time-course experiments showed that the presence of CBM5 enhanced early hydrolysis on crab shell chitin (Fig. 1B), suggesting that the domain increases the reaction rate during initial substrate hydrolysis. A similar trend was observed for chitinolytic activity on chitin azure (CA) (Fig. 1C, D) but not for activity on >75% de-acetylated chitosan, where SiCHIT and SiCHIT−CBM5 displayed similar reaction rates already at the onset of the reaction (Fig. 1E).

To evaluate whether CBM5 influences product outcome, we analyzed the hydrolysates of crab shell chitin by mass spectrometry. Both SiCHIT and SiCHIT−CBM5 produced similar oligomer profiles dominated by fully or partially acetylated dimers (A2 or A1D1), indicating that CBM5 does not alter product specificity (Fig. 1F, Supplementary Fig. 2). Notably, substantial amounts of other partially deacetylated products, such as A1D2 and A2D1, were also detected. This likely reflects the fact that crab shell chitin is not fully acetylated; deacetylated regions may form stretches with higher solubility and increased accessibility, leading to their preferential cleavage by the enzymes. Our mass spectrometric analysis revealed a residual activity of SiCHITE196Q that was not detectable with the other, less sensitive activity assays using crab shell chitin or CA. We also analyzed product profiles on β-chitin (Supplementary Fig. 3). While truncation of the CBM5 domain did not affect the product profile on this substrate, slight differences were observed depending on the substrate. These differences can be attributed to the less rigid structures of β-chitin, which result from its more loosely packed polymer folding.

To assess whether CBM5 affects subsite preferences, we analyzed the hydrolysis products of chitosans with defined degrees of acetylation (DA). SiCHIT−CBM5 displayed the same requirement for an acetylated unit at the −1 subsite and no detectable shifts at adjacent positions compared to SiCHIT31, confirming that CBM5 does not influence catalytic specificity (Fig. 1G, Supplementary Fig. 4).

To determine the role of CBM5 in substrate interaction, we performed pull-down assays using crab shell chitin, >75% de-acetylated chitosan, and lyophilized, washed Bs mycelium. SiCHIT−CBM5 showed reduced binding to crab shell chitin compared to full-length SiCHIT, whereas no difference was observed with chitosan, suggesting that CBM5 enhances binding specifically to acetylated, crystalline substrates (Fig. 1H; Supplementary Fig. 5). This is consistent with our observation that CBM5 increases the initial degradation rate on crab shell chitin but not on chitosan (Fig. 1B, E). Binding to Bs mycelium was likewise reduced in the absence of CBM5, indicating that the domain contributes to effective fungal cell wall targeting (Fig. 1H).

Together, these results show that although CBM5 is dispensable for full catalytic activity and does not alter product specificity, it enhances early hydrolysis rates on chitin and increases substrate binding to crystalline chitin and fungal mycelium. This suggests that CBM5 improves functional efficiency under biologically relevant conditions, likely contributing to the role of SiCHIT in microbial competition.

CBM5 is required for antifungal activity and plant protection
We previously demonstrated that SiCHIT restricts the growth of Bs and reduces its colonization of plant roots through its chitinolytic activity31. However, the specific contribution of the CBM5 domain to this antifungal function remained unresolved. To test whether CBM5 is required for Bs inhibition, we incubated Bs spores with the full-length SiCHIT, the catalytically inactive mutant SiCHITE196Q, the truncated variant lacking CBM5 SiCHIT−CBM5, or the empty vector control Ev. As expected, Bs spore germination was significantly reduced in the presence of active SiCHIT, but not with SiCHITE196Q. SiCHIT−CBM5 showed a similar lack of inhibitory effect as SiCHITE196Q and had only a minimal effect compared to the Ev control (Fig. 2A).

We next examined whether the loss of antifungal activity translates into reduced protection of host roots. Bs spores treated with the same chitinase variants were used to inoculate the roots of the Bs host Hordeum vulgare (Hv). As previously shown, SiCHIT-treated spores led to a significant reduction in Bs colonization, as measured by fungal transcript levels in roots. In contrast, SiCHITE196Q and SiCHIT−CBM5 had no effect (Fig. 2B). Correspondingly, only SiCHIT mitigated the reduction in root biomass caused by Bs infection (Fig. 2C). To test whether this effect is conserved in other hosts, we repeated the experiment using Arabidopsis thaliana (At). SiCHIT reduced Bs biomass and preserved root elongation, while SiCHITE196Q did not. SiCHIT−CBM5 had a weak protective effect, albeit significantly lower than the full-length enzyme (Supplementary Fig. 6).

Together, these results demonstrate that both chitinase activity and the CBM5 domain are required for SiCHIT’s full antifungal and plant-protective functions. The loss of protection observed in the CBM5-lacking variant indicates that this domain enhances fungal cell wall engagement and is critical for effective niche defense.

A CBM5-lacking paralog, SiCHIT2, is transcriptionally specialized for host colonization
GH18-CBM5 chitinases are absent in Ascomycota but occur frequently in the Agaricomycetes (Basidiomycota), where gene copy number varies across species with different ecological lifestyles31. To better define the structural requirements for antifungal activity in Si, we compared the four GH18 chitinases encoded in the genome with respect to phylogenetic relationships, domain architecture, and transcriptional regulation. Of the four, only SiCHIT harbors a CBM5 domain, whereas the other three chitinases contain no additional domains besides the GH18 catalytic domain (Fig. 3A). SiCHIT and its closest paralog SiCHIT2 (PIIN_03542; Pirin1_74345) share the highest sequence identity (69.3%) and form a distinct phylogenetic pair among the Si GH18 chitinases (Fig. 3A; Supplementary Fig. 7). The catalytic DxxDxDxE motif located at the −1 subsite is conserved across all four chitinases, suggesting that each is catalytically active (Fig. 3B). Despite this shared motif, the chitinases exhibit distinct structural features. SiCHIT2 retains an 86-amino-acid C-terminal interdomain sequence (linker) but lacks a CBM5 domain. SiCHIT3 (PIIN_11727; Pirin1_71855) lacks a predicted signal peptide, while SiCHIT4 (PIIN_07603; Pirin1_78411) features extended surface-exposed loops within the GH18 domain (Supplementary Table 1).

The evolutionary relationships among fungal GH18 chitinases, reconstructed using maximum-likelihood phylogenies of the catalytic domain (Supplementary Figs. 8 and 9), reveal three major fungal clades, including one enriched in CBM5 domain-containing (GH18–CBM5) chitinases. SiCHIT and SiCHIT2, which are tandem, adjacent genes in the Si genome, form a close sequence and structural pair distinct from other Si chitinases (Fig. 3C and Supplementary Figs. 8 and 9). Their high sequence identity and the absence of CBM5 in SiCHIT2 are consistent with a recent gene-duplication event followed by domain loss in one paralog. This supports a model in which the two genes may have undergone neo-functionalization or sub-functionalization within Si.

Across the broader fungal phylogeny, the GH18–CBM5-rich clade includes species with numerous paralogs, some of which also lack CBM5, consistent with multiple independent domain-loss events across Basidiomycota, or with the possibility that these patterns arose through tandem duplication followed by domain loss in an ancestral lineage and subsequent vertical inheritance. The observed lack of CBM5 is not uniquely associated with plant-associated lineages, indicating that domain loss is likely driven by diverse ecological pressures and functional requirements rather than lifestyle alone. The presence of the GH18–CBM5 domain combination in bacterial sequences that form an outgroup to this clade, together with the apparent absence of GH18–CBM5 chitinases in Ascomycota, raises the possibility that the CBM5-bearing GH18 lineage in Basidiomycota may have originated through horizontal gene transfer from bacteria, representing one plausible evolutionary scenario (Supplementary Figs. 8 and 9). Overall, this phylogenetic pattern supports a complex evolutionary history for GH18 chitinases involving gene duplication, domain gain and loss, and functional diversification.

To assess structural similarity between SiCHIT and SiCHIT2, we generated AlphaFold models42,43 of both enzymes after removing the predicted signal peptides. The models confirmed the overall structural conservation of the GH18 catalytic domain and the presence of a C-terminal linker in SiCHIT2, but no CBM5 domain (Fig. 3D and E). The GH18 with the catalytic DxxDxDxE motif could also be confirmed in SiCHIT3 and SiCHIT4, with the latter displaying three additional loop structures (Supplementary Fig. 10A, B).

To examine transcriptional regulation, we analyzed RNA-seq data from Eichfeld et al.31 and found that SiCHIT and SiCHIT2 were the only chitinases significantly upregulated in biotic contexts (Fig. 3F). SiCHIT was specifically induced during fungal confrontation, while SiCHIT2 was induced during colonization of barley, Brachypodium, and Arabidopsis. In contrast, SiCHIT3 and SiCHIT4 were not transcriptionally activated under any tested biotic conditions, including interactions with pathogenic fungi, beneficial bacteria, or different plant hosts (Fig. 3F, G). To extend the RNA-seq findings, we quantified the expression of all four GH18 chitinases in Si by qPCR following confrontation with Bs and six additional filamentous fungi representing the major fungal lineages Ascomycota, Basidiomycota, and Mucoromycota. SiCHIT was induced in several of these interactions, whereas expression of SiCHIT2, SiCHIT3, and SiCHIT4 remained low under all tested microbial conditions (Fig. 3H). When Si was grown on crab shell chitin as the sole carbon source, SiCHIT, SiCHIT3, and SiCHIT4 were expressed at varying levels, whereas SiCHIT2 expression remained comparatively low (Fig. 3I). Because these paralogs show distinct transcriptional profiles, we hypothesized that diversification in their upstream regulatory regions may underlie these differences. We therefore compared up to 500 bp upstream of each coding sequence using MEME Suite44. As a reference, we analyzed the promoter regions of six DELD effector genes, a known expanded gene family in Si45,46. Whereas DELD promoters displayed strong motif conservation, the putative regulatory regions of the four chitinases were highly divergent (Supplementary Fig. 11). Consistent with this divergence, weighted correlation network analysis (WCNA) shows that the four Si chitinases fall into distinct transcriptional modules: SiCHIT clusters in module 6, SiCHIT2 in module 4, SiCHIT3 in module 1, and SiCHIT4 in the unassigned background module 0 (Supplementary Fig. 1A and Supplementary Table 1 in Brands et al.47). Notably, module 4 is plant-responsive and enriched for secreted proteins, including multiple predicted effectors such as DELDs, SiWSC3, SiFGB1, SiE5NT, and SiNucA, indicating that SiCHIT2 is co-expressed with a host-induced secretome program.

Together, these findings suggest that SiCHIT2 evolved a distinct function from SiCHIT following gene duplication. Their divergent domain architecture and transcriptional profiles are consistent with functional specialization, with SiCHIT adapted to fungal antagonism and SiCHIT2 to host-associated functions during colonization.

Fusion of CBM5 reprograms SiCHIT2 for fungal antagonism
To test whether the CBM5 domain of SiCHIT is sufficient to confer antifungal activity, we constructed a chimeric protein consisting of the GH18 catalytic domain of SiCHIT2 fused to the proline-rich linker and CBM5 domain of SiCHIT (SiCHIT2+CBM5). As a control, we generated a second construct in which only the linker was fused to the GH18 domain of SiCHIT2 (SiCHIT2+link) (Fig. 4A). As observed for native SiCHIT and SiCHIT−CBM5, the presence of CBM5 did not affect endpoint chitinase activity on crab shell chitin or chitin azure (Fig. 4B, C) but enhanced early hydrolysis (Supplementary Fig. 12). Notably, approximate Michaelis–Menten kinetics revealed that the presence of CBM5 reduced the maximal reaction velocity (Vmax) while decreasing Km (Supplementary Fig. 13), indicating increased substrate affinity but a modest constraint on catalytic turnover under saturating substrate conditions. Together with the time-course assays on crab shell chitin and chitin azure (Fig. 1B, D; Supplementary Fig. 12), these findings indicate that CBM5 promotes substrate degradation during early reaction stages, while imposing steric limitations at substrate saturation. However, only SiCHIT2+CBM5 negatively affected Bs spores, comparable to the effect of full-length SiCHIT, whereas SiCHIT2+link showed no inhibitory activity (Fig. 4D). This pattern was reflected in the hydrolytic activity on lyophilized Bs mycelium, where SiCHIT and SiCHIT2+CBM5 showed strong activity (Supplementary Fig. 14A, B). Notably, SiCHIT−CBM5 and SiCHIT2+link also displayed low activity on Bs mycelium. Chitinases lacking CBM5 may partially degrade the cell wall but appear unable to further hydrolyze cell wall fragments, as indicated by the non-migrating material observed in the TLC assay (Supplementary Fig. 14B). Consistent with these results, both SiCHIT and SiCHIT2+CBM5 alleviated Bs-induced root growth inhibition in barley and reduced pathogen colonization, whereas SiCHIT−CBM5 and SiCHIT2+link had no measurable protective effect (Fig. 4E, F).

These results demonstrate that fusing CBM5 to SiCHIT2 is sufficient to confer antifungal activity, indicating that CBM5 acts as a modular domain enabling functional adaptation toward microbial antagonism.

SiCHIT2 suppresses chitin-triggered immunity and facilitates root colonization by S. indica
Given the distinct plant-induced expression profile of SiCHIT2, we hypothesized that this chitinase contributes to immune evasion during root colonization. Previous studies have shown that Podosphaera xanthii uses chitinase-like effectors to degrade immunogenic chitin oligomers and suppress host defense responses48. To test whether Si chitinases function similarly, we incubated immunogenic chitohexaose (A6) with recombinant SiCHIT, SiCHIT2, their respective variants, or the Ev control. The resulting hydrolysates were applied to barley roots to measure reactive oxygen species (ROS) production. ROS accumulation was completely abolished when chitohexaose was pre-treated with either SiCHIT2+link or SiCHIT−CBM5. In contrast, treatment with SiCHIT2+CBM5 or full-length SiCHIT resulted in only partial or no suppression of ROS accumulation (Fig. 5A, B). An intermediate reduction in ROS accumulation was observed when chitohexaose was pre-treated with SiCHITE196Q, likely due to the inactive catalytic cleft sequestering a fraction of the chitohexaose. None of the recombinant chitinases induced ROS when applied in the absence of chitohexaose (Supplementary Fig. 15).

To assess whether immune evasion contributes to host colonization, we co-inoculated Si with 20 µM recombinant chitinase proteins and quantified fungal biomass in barley roots 3 days post-inoculation. SiCHIT−CBM5 and SiCHIT2+link significantly enhanced root colonization, whereas the other chitinases did not (Fig. 5C). In bacterial GH19 and GH18 chitinases, removal of CBM5 has been shown to accelerate the degradation of small, soluble chitooligomers40,49, likely by reducing steric hindrance at the catalytic center. We therefore hypothesized that differences in the ability to evade chitin-triggered immunity might reflect differences in the degradation of soluble immunogenic chitooligosaccharides. To test this, we measured the chitinolytic activity of all recombinant variants on chitohexaose using a DNSA reducing-end assay. SiCHIT−CBM5 and SiCHIT2+link showed higher release of reducing ends from chitohexaose compared to SiCHIT or SiCHIT2+CBM5 (Supplementary Fig. 16A, B). In addition, time-resolved SEC-RI-MS analysis indicated that recombinant chitinase protein variants lacking the CBM5 domain depleted chitohexaose and the intermediate chitotetraose more rapidly, resulting in faster accumulation of shorter oligomers (DP2-3) (Supplementary Fig. 16C, D). Because chitin-triggered ROS production is both threshold- and time-dependent, this kinetic bias provides a parsimonious explanation for the stronger suppression of immunity observed for these variants; however, additional in planta factors (e.g., protein stability/availability and solubility in the apoplast) may further modulate the magnitude of the response.

Expression of the barley defense gene PR10 was similar across treatments at 3 dpi despite increased colonization by SiCHIT−CBM5 and SiCHIT2+link (Fig. 5D). This suggests that immune evasion mediated by the recombinant SiCHIT−CBM5 and SiCHIT2+link occurs primarily early during host contact, facilitating initial colonization. These results suggest that SiCHIT2’s ability to evade ROS generation depends on the absence of CBM5, which may otherwise restrict access to soluble substrates and favor binding to insoluble chitin. This immune-evasive function, together with its host-specific transcriptional induction, underscores a distinct role for SiCHIT2 compared to its CBM5-containing paralog.

Methods
Plant and fungal materials
Hordeum vulgare (Hv, L. cv Golden Promise) and Arabidopsis thaliana (At, Col-0) were used as plant hosts. Hv seeds were surface-sterilized in 6% sodium hypochlorite for 1h under constant shaking and subsequently washed five times for 30min in sterile water. Seeds were germinated for 4 days on wet filter paper at 21°C in the dark. Seeds of At were surface-sterilized in 70% ethanol for 10min and 100% ethanol for 7min. At seeds were placed on ½ MS medium (Murashige-Skoog medium including vitamins, pH 5.6) containing 1% sucrose and stratified for 3 days in the dark and 4°C prior to germination on a day–night cycle of 8/16h at 22/18°C, 60% humidity, and 125µmol/m2 light intensity for 7 days. Serendipita indica (Si, DSM11827) and Bipolaris sorokiniana (Bs, ND90Pr) were used as fungal species. Si was grown on CM medium (Complete Medium, 1.5% agar61) at 28°C for 28 days in the dark. Si spores were harvested by adding sterile water and scraping the spores with a scalpel. The suspension was filtered through a sterile Miracloth filter and spores were centrifuged at 2000×g. The supernatant was discarded and spores were washed three times in water. Bs was grown on modified CM medium25 at 28°C for 14 days in the dark. Bs spores were harvested like Si spores, but a Drigalski spatula was used to scrape the spores instead of a scalpel.

Preparation of fungal inoculants and plant colonization
For the preparation of Bs inoculants, spores were harvested from 21-day-old plates and incubated with the respective recombinant proteins (10µM in 50mM phosphate buffer) for 16h at 28°C in the dark. For the preparation of Si inoculants, spores were harvested from 28-day-old plates and allowed to germinate overnight in sterile water at 28°C. Prior to plant root inoculation, the respective recombinant proteins were added to 20µM in 50mM phosphate buffer. Spore mixtures were diluted to 5000 spores/mL for Bs and 500,000 spores/mL for Si to yield the final working inoculants. For inoculation of At, ten sterile 7-day-old seedlings were placed on solid 1/10 PNM (pH 5.6) plates (0.005% KNO3, 0.005% KH2PO4, 0.0025% K2HPO4, 0.049% MgSO4, 0.00472% Ca(NO3)2, 0.0025% NaCl, 0.5% (v/v) Fe-EDTA stock solution, 1.2% Gelrite, pH 5.6. After autoclaving, 10mL 1M MES pH 5.6 was added. The EDTA stock solution contains 2.78% FeSO4×7H2O, 4.13% Na2EDTA×2H2O). Concentrations are indicated in w/v if not stated otherwise. The roots were treated with 1mL of the respective inoculant and grown on a day–night cycle of 8/16h at 22/18°C, 60% humidity and 125µmol/m2 light intensity for 3 days. The root elongation was quantified by measuring the difference in root length at the start of the inoculation and the end of the experiment at 3 days post-inoculation (dpi). Roots were washed with water and snap frozen in liquid nitrogen for further RNA extraction. For inoculation of Hv, four sterile seedlings were placed in glass jars containing solid 1/10 PNM medium (pH 5.6) and inoculated with 3mL of the respective inoculant. The inoculated plants were grown on a day–night cycle of 16/8h at 22/18°C and 60% humidity and a light intensity of 108µmol/m2 for 3 days. Root weight was quantified at 3 dpi. Roots were washed in water and snap frozen for further RNA extraction.

Preparation of fungal confrontations
For the preparation of fungal inoculants, liquid CM medium was inoculated with 1 mL of S. indica spores at a concentration of 500,000 spores/mL or 1 mL of B. sorokiniana spores at a concentration of 5000 spores/mL. For all other fungi, mycelium was scraped from plates and transferred to liquid CM. All fungi were grown for 6 days at 28°C and 120rpm. The mycelium was blended and re-suspended in fresh CM61 and regenerated for another 24h. Regenerated mycelium was filtered through Miracloth filters and washed thoroughly with sterile MilliQ water. Fungal mycelia were mixed in a 1:1 ratio, and 1g of mycelium was streaked on PNM plates containing 15% agar. After 6 days, the mycelia were harvested, dried with tissue papers, and snap frozen in liquid nitrogen.

Antifungal activity assay
To quantify the Bs spore germination, isolated spores were resolved in liquid CM to a concentration of 100,000 spores/mL and 100µL were added to the wells of sterile Lab-Tek 2 chamber slides with cover (VWR). 100µL of the respective treatment (Ev control or purified protein) was added to a final concentration of 10µM. The germination rate of spores was quantified 6h after treatment.

RNA extraction from plant roots and qPCR
Plant materials were ground in liquid nitrogen, and 1mL of TRIzol reagent (Thermo Fischer Scientific) was added. Samples were vortexed for 20s until all material was resolved. 200µL chloroform was added, and samples were vortexed again. Samples were centrifuged at 17,000×g, and 4°C for 30min and 500µL of the upper aqueous phase was transferred to 500µL isopropanol. The nucleic acids were precipitated at −20°C for 16h and pelleted by centrifugation at 17,000×g and 4°C for 30min. Pellets were washed three times with 70% ethanol diluted in DEPC water. Remaining DNA was removed by adding 1µL DNase 1 (1U/µL) in 3µL DNase 1 buffer (10x) (Thermo Fischer Scientific) and 26µL RNase-free water. Samples were incubated at 37°C for 30min. DNase digest was stopped by heating to 70°C for 5min and subsequent precipitation with 2 volumes of isopropanol and ½ volume of 7.5M ammonium acetate for 16h at −20°C. Samples were washed twice with 70% ethanol in DEPC water and resolved in 30µL RNase-free water. Concentrations were determined using a NanoDrop 2000c spectrophotometer (Thermo Fischer Scientific).

1µg of RNA was used for reverse transcription into cDNA (Thermo Fischer Scientific). First, 1µL of random hexamer and of oligo-dt oligomers were added and incubated at 65°C for 5min and subsequently placed on ice. 4µL 5× reaction buffer, 1µL Riboblock RNase inhibitor (20U/µL), 2µL 10mM dNTPs, and 2µL MMLV reverse transcriptase were added, and samples were incubated at 42°C for 1h. cDNA synthesis was stopped by heating to 70°C for 5min. qPCR was conducted using the primers listed in Supplementary Table S2.

Cloning of chitinase expression vectors
The coding sequence (CDS) of the chitinase genes was amplified by PCR with Q5 High-Fidelity Polymerase (New England Biolabs) using the primers listed in Supplementary Table 2. PCR products were cloned using the Gibson assembly cloning technique into the PQE80L expression vector (Qiagen, Hilden, Germany) after vector linearization with Kpn1 (New England Biolabs). For the production of SiCHIT2+link and SiCHIT2+CBM5, the inter-domain region (linker) or the inter-domain region with the CBM5 domain of SiCHIT2 was fused to the C-terminus of the GH18 domain of SiCHIT2.

Production and purification of recombinant protein in E. coli
Chemically competent E. coli BL21 cells were transformed with the vectors for protein expression under control of a lac operon. 5mL starter cultures were prepared in liquid LB medium containing 100µg/mL carbenicillin and grown for 16h at 37°C and 180rpm. Starter cultures were transferred to 300mL of LB medium with carbenicillin and grown to an OD600 of 0.5 at 28°C. Gene expression was induced by adding IPTG to a final concentration of 1mM and cells were grown at 16°C for 20h. Cells were pelleted by centrifugation at 5000×g for 15min at 4°C. Bacterial pellets were resolved in 5mL lysis buffer (50mM NaH2PO4, 300mM NaCl, and 10mM imidazole, pH 8) and sonicated three times for 1min with pulses on ice and centrifuged at 16,200×g for 30min. 5mL of the supernatant was added to 1mL of Nickel-NTA Agarose (Qiagen, Hilden, Germany) and incubated under continuous shaking at 4°C for 1h. Samples were spun down at 500×g for 10s, the flow through was removed, 4mL wash buffer (50mM NaH2PO4, 300mM NaCl, and 20mM imidazole, pH 8) was added, and samples were incubated under continuous shaking at 4°C for 10min. Washing steps were repeated five times. For elution, 1mL of elution buffer (50mM NaH2PO4, 300mM NaCl, and 250mM imidazole, pH 8) was added, and samples were incubated under continuous shaking at 4°C for 10min. Eluted fractions were dialyzed against 50mM phosphate buffer (pH 6) overnight, and protein concentrations were measured using Bradford reagent. To test the purity of the protein samples, aliquots were boiled for 5min in SDS-sample buffer, loaded on a 10% SDS gel, and visualized by Coomassie staining.

Chitinase activity assays
To quantify the enzymatic activity of the purified chitinases, 2mg/mL chitin azure (Sigma Aldrich; C3020) was incubated with 5µM recombinant chitinase in a total volume of 200µL. Reactions were incubated at 28°C for 20h. Reactions were stopped by heating to 95°C for 5min, and samples were centrifuged 5min at 17,000×g and the absorbance of supernatants was quantified at 560nm. For quantifying reducing ends, 20mg crab shell chitin (Sigma Aldrich; C7170) or >75% de-acetylated chitosan from shrimp cells (Sigma Aldrich; C3646, average molecular weight ~360kDa) were incubated with 10µM enzyme in 200µL 50mM phosphate buffer for 20h at 28°C. Reactions were stopped by heating to 95°C for 5min. Samples were centrifuged for 5min at 17,000×g, and the supernatant was mixed in a 1:1 ratio with DNSA (3,5-dinitrosalicylic acid) reagent. DNSA reagent is prepared by mixing 1% (w/v) 3,5-dinitroslicylic acid, 1.6% (w/v) sodium hydroxide, 30% (w/v) potassium sodium tartrate tetrahydrate. The samples were heated for 10min at 95°C and subsequently transferred to ice. The absorbance of the samples was measured at 540nm. OD540 values were converted to the concentration of reducing ends (mol/L) using a N-acetylglucosamin standard series.

For time course experiments with CA, crab shell chitin and chitosan, samples were prepared as described above in a final volume of 1mL and placed in a 28°C incubator, allowing reaction tubes to shake horizontally. 150µL of the samples was taken every hour.

For the kinetic analysis, crab shell chitin at concentrations from 10 to 120mg/mL was incubated with 5µM recombinant enzyme for 60min. Curve fitting and Michaelis–Menten parameters were calculated using the package “renz” and “drc” in R.

For TLC on Bs mycelium, 15mg lyophilized and washed mycelium were weighed in 2mL reaction tubes. For washing, lyophilized mycelium was heated at 80°C for 5min in 1mL 50mM phosphate buffer and then washed in 50mM phosphate buffer until supernatants did not color Bradford dye (5 times). The recombinant chitinases were added to a final concentration of 5µM in 200µL. Reactions were incubated at 28°C for 20h and stopped by heating to 95°C for 5min. Samples were centrifuged at 17,000×g for 5min. 10µL of the supernatant was separated on 60 F254 TLC (Sigma Aldrich) silica gel plates with running buffer (n-butanol/methanol/28% ammonium hydroxide/water, 5/4/2/1). Plates were dried and then bathed in developer solution (1.6g diphenylamine/1.6mL aniline/12mL 85% phosphoric acid/80mL acetone). TLC plates were developed in an incubation oven at 100°C for 20–30min.

To determine product profiles, 1mg of each insoluble substrate was incubated with the corresponding enzyme (5µM) in 40µL of ammonium acetate buffer (50mM, pH 5) for four days at 37°C and 180rpm. Subsequently, the samples were filtered through centrifugal filters (modified PES, 3kDa cutoff; VWR, Germany), dried in a vacuum concentrator at 30°C, and resuspended in 10µL MilliQ water. A volume of 2µL was analyzed using the HILIC-ESI-MS1 system described above in sensitivity mode. Different from subsite specificity analyses, the column temperature was set to 35°C, and the following gradient elution profile was used: 100% A for 0.0–2.0 min, followed by a linear gradient to 75% (v/v) B from 2.0 to 5.0 min. From 5.0 to 8.0 min, the flow was isocratic 75% (v/v) B, followed by a linear gradient from 25% (v/v) A to 100% A from 8.0 to 8.2 min, and an isocratic elution of 100% A from 8.2 to 10.0 min. Resulting data was analyzed using MassLynx V4.2 (Waters, USA) and the intensities of the MS signals for each oligomer were quantified using Skyline 23.1 (MacCoss Lab Software, University of Washington, USA63) as described above, exemplary mass spectra are shown in Supplementary Fig. 2.

Chitinase subsite specificity and product profiles
The subsite specificity of SiCHIT−CBM5 was determined as previously reported31,62 in biological triplicates. SiCHIT-CBM5 was used at a final concentration of 0.2µM on 1mg/mL of chitosans with an average degree of acetylation (DA) of 10%, 35%, or 50% and an average degree of polymerization (DP) around 1000 in 50mM of sodium acetate buffer. Incubation was performed for 1h or 24h. Oligomers produced by SiCHIT-CBM5 were N-acetylated using [2H6]acetic anhydride, and MS1 samples were subsequently mixed with internal chitin standards with DPs from 1-6, which are double-isotopically labeled via N-acetylation with [2H6;13C4]acetic anhydride. These were analyzed by HILIC-ESI-MS1 using a Waters ACQUITY Premier UPLC System coupled to a Waters Synapt XS HDMS 4k mass spectrometer. Oligomer mixtures were separated using an Acquity UPLC BEH Amide column (1.7μm, 2.1mm×50mm; Waters Corporation, Milford, MA, USA) and a VanGuard precolumn (1.7μm, 2.1mm×5mm; Waters Corporation, Milford, MA, USA). The flow rate was set to 0.4mL/min, the column oven temperature was set to 40°C, and 2µL of the sample, including 50ng of each of the internal standards (double isotopic labeled chitin oligomers of DP 1–6), was injected into the system. The sample was separated using an LC run lasting 10.5min, with the following gradient elution profile: 100% A (80:20 acetonitrile:H2O with 10mM NH4HCO3 and 0.1% (v/v) HCOOH) for 0.0−1.0 min, followed by a linear gradient to 20% (v/v) B (20:80 acetonitrile:H2O with 10mM NH4HCO3 and 0.1% (v/v) HCOOH) from 1.0-7.5 min, and then to 75% (v/v) B from 7.5 to 8.5 min. From 8.5 to 9.0 min, the flow was isocratic 75% (v/v) B, followed by a linear gradient from 25% (v/v) A to 100% A from 9.0 to 9.2 min, and an isocratic elution of 100% A from 9.2 to 10.5 min. The MS1 measurements were conducted in positive resolution mode with normal dynamic range sensitivity, with a mass range of m/z 50–2000 and a scan time of 0.25s. To improve mass accuracy, lock spray calibration was performed using leucine encephalin, injected in 10-s intervals for one second as a lock mass. The capillary voltage of the source was set to 3kV, the sampling cone to 35, the source offset to 20, while the source and desolvation temperatures were set to 80 and 250°C, respectively. Cone gas flow was set to 0 l/h, and the desolvation gas flow was set to 500 l/h. The nebulizer pressure was set to 5.8 bar.

Quantitative pattern determination by MS2 was performed following the method previously described62. Briefly, the oligomers produced by SiCHIT−CBM5 after N-acetylation using [2H6]acetic anhydride as described above were incubated in H218O overnight to label the reducing end of the products with 18O before MS2 analysis. The Waters LC–MS system used for the MS1 analysis was also used for this analysis. The TOF MS/MS mode was employed to determine the pattern of the N-acetylated and 18O-labeled oligomers. Each oligomer containing at least one natural and one isotopically labeled acetyl group was individually isolated using the quadrupole of the MS system with an LM resolution of 15 to enable the isolation and fragmentation of monoisotopic peaks. The collision energy in the trap was set to 18V for dimers, 20V for trimers, 22V for tetramers, 24V for pentamers, and 26V for hexameric products. MS data was analyzed using MassLynx V4.2 (Waters, USA) and the intensities of the MS signals for each oligomer or fragment were quantified using Skyline 23.1 (MacCoss Lab Software, University of Washington, USA63). Exemplary mass spectra are shown in Supplementary Fig. 4. The MS1 signal intensities of each product were compared to the intensity of the corresponding double isotopic-labeled standard to calculate the amount of each product based on the 50ng of injected standard. The MS2 signal intensities of a-fragments and the largest b-fragment were used to calculate the proportions of the different patterns of the oligomers64. The combination of the resulting datasets allows for calculating the amount of each product of defined DP, DA, and pattern, which can in turn be used to quantify the frequency of acetylated units at the −2 to +2 subsites of SiCHIT−CBM5.

SEC–RI–MS time-resolved product profiling on chitohexaose
Product profiles generated by the five enzymes during hydrolysis of the chitin hexamer (A6) were monitored by live sampling of ongoing reactions coupled to size exclusion chromatography with refractive index and mass spectrometric detection (SEC–RI–MS), as described previously65. Briefly, A6 (0.5mg/mL) was prepared in ammonium acetate buffer (150mM, pH 4.5 adjusted with acetic acid) and pre-heated to 37°C. Reactions were assembled directly in the autosampler by mixing 20µL substrate solution with 10µL preheated enzyme stock (0.3µM in the same buffer), yielding final concentrations of 0.33mg/mL A6 and 0.10µM enzyme in a total volume of 30µL. Immediately after mixing, 3µL were withdrawn and injected as the first timepoint (1.5min). Subsequent 3µL injections were taken at 17.0, 32.5, and 48.0min (i.e., every 15.5min after the initial 1.5min timepoint) while incubating at 37°C in the autosampler. MS and RI data were exported in DataAnalysis 4.1 (Bruker) and processed in Skyline 23.1 (MacCoss Lab Software)63 and OriginPro 2025 (OriginLab), respectively.

Substrate binding assay
15mg of crab shell chitin (Sigma Aldrich; C7170), Chitosan (Sigma Aldrich; C3646) or lyophilized mycelium of Bs were weighed in a 2mL reaction tube. For washing, lyophilized mycelium was heated at 80°C for 5min in 1mL 50mM phosphate buffer and then washed in 50mM phosphate buffer until supernatants did not color Bradford dye (5 times). Samples were incubated with 400µL chitin-binding buffer (2mM KH2PO4, 8mM Na2HPO4, 2mM KCL) and 100µL of 20µM recombinant protein, shaking at 4°C for 1h. Samples were centrifuged at 17,000×g for 5min and 60µL of the supernatant was mixed with 60µL Bradford reagent. Protein amounts were quantified at 595nm and compared to the input amount. In addition, supernatants or pellets were boiled in SDS-sample buffer at 95°C and separated via SDS–PAGE on a 10% SDS gel and stained with Coomassie brilliant blue.

Quantification of reactive oxygen species (ROS)
For elicitor preparation, 10µM of chitohexaose was incubated with 10µM of recombinant chitinase or the buffer control for 20h at 28°C. Reactions were stopped by heating to 95°C for 5min and chitohexaose was diluted to 250nM.

Four root pieces (0.5cm) of 4-day-old barley seedlings were transferred to each well of a 96-well microtiter plate (white, flat bottom) containing 200μL of 2.5mM 2-(N-morpholino)ethanesulfonic acid (MES) buffer, pH 5.6. The plate was incubated for 16h at 21°C for recovery. The buffer was replaced with 100μL 2.5mM MES buffer containing 20μM LO-12 (Wako Chemicals) and 20μg/mL Horseradish peroxidase (HRP) (Sigma-Aldrich). After 25min incubation in the dark, 100μL of 250nM chitohexaose elicitor solution was added to each well. Chemiluminescence was measured using a TECAN SPARK 10M microplate reader over all wells for 2h with an integration time of 450ms.

Fungal sequence extraction and GH18 domain extraction
Fungal sequences were manually extracted from JGI Mycocosm portal66 (Supplementary File 1) and only species encoding GH18-CBM5 were included. Basidiomycetes and Ascomycetes were surveyed for GH18-CBM5 sequences, and no hits were found for Ascomycetes (see below). Therefore, GH18 sequences from Ascomycetes were excluded from further analysis. In total, 985 fungal GH18 sequences, 212 of which contained CBM5 motifs, were extracted. From the GH18 domains of the Si and Sv chitinases, a Hidden Markov model (HMM) profile was built using the hmmbuild function from the hmmer67 package (version 3.3.2) in the Linux terminal. This GH18 hmm profile was used as input for the hmmsearch function from the same package to search against the 985 downloaded fungal sequences. From the hmmsearch results, sequences with a best domain score higher than 200 were selected. In total, 240 fungal sequences passed this threshold, 51 of which had the CBM5 motif. All sequences belonged to the phylum Basidiomycota. The GH18 domains of these sequences were extracted using the same hmmsearch function with the –domtblout argument, obtaining the starting and ending position of the GH18 HMM domain per sequence from the envelope columns. For sequences that had multiple GH18 domains, the GH18 domain closest to the C-terminal was chosen. Only the GH18 domain of these sequences was used in the subsequent analysis (Supplementary File 1).

Bacterial sequence extraction
Bacterial GH18 sequences were extracted from UniprotKB (release 2025_04) using the following search terms: (taxonomy_id:2) AND (xref:pfam-PF00704). Taxonomy ID 2 refers to bacteria and PF00704 refers to the Pfam annotation for the GH18 motif. The hmmsearch of the Si/Sv GH18 hmm profile was performed against the bacterial sequences, similarly to what was done before the extraction of the GH18 domain. Based on this, two types of outgroup sequences were chosen. The first group contained bacterial sequences that had a best domain score of more than 200. 5 GH18 sequences were picked for this group, corresponding to 3 bacterial species that were reported to live in soil and/or colonized plants. For two of the three species, two sequences were chosen, one from a CBM5-containing sequence and one without. From the third species, only the CBM5-containing sequence passed the threshold. The second bacterial group contained GH18 sequences with a low best domain score (between 20 and 30) from the hmmsearch results. 5 random sequences were chosen from that group, and they were from non-CBM5-containing sequences.

Sequence-based distinction between CBM5 and CBM12
In the InterPro and Pfam annotations, CBM5 and CBM12 domain sequences are not distinguished. To clearly separate the CBM5 motif and the closely related CBM12 motif, CBM5 and CBM12 sequences were downloaded from the CAZy68 database. Then, sequences found in both entries were discarded to remove any potential overlap. 20 random sequences per the remaining CBM5 or CBM12-specific sequences were chosen to build a respective HMM profile using the hmmbuild from hmmer package (version 3.3.2) (Supplementary Fig. 17). Next, an hmmsearch using either the CBM5 or CBM12 hmm profile was executed against either CBM5 or CBM12-specific sequences that were not included in building the hmm profile. For the CBM5 sequences, the best domain score from the hmmsearch of both the CBM5 and CBM12 motifs was compared. The same was done for the CBM12 sequences. For CBM12-specific sequences, the CBM12 hmm score was higher than the CBM5 score (n=162; Wilcoxon test; p-value < 2.2e–16; V=13162). For CBM5-specific sequences, the corresponding CBM5 hmm score was higher (n=164, Wilcoxon test; p-value < 2.2e−16; V=13108).

To confirm that the CBM5 sequences used were truly CBM5 and not CBM12, the CBM5 and CBM12 hmm search was applied on them, which confirmed that the CBM5 hmm score is higher than the CBM12 one.

Sequence processing and phylogenetic inference
GH18 sequences were aligned using MAAFT69 (version 7). After the alignment, the sequences were trimmed to remove uninformative sites using Clipkit70 (Version 2.6.1) on the Linux terminal. Next, the sequences were used as input for IQ-TREE71 (version 2.0.7) on the Linux terminal. Model inference without tree reconstruction was conducted using the -MF argument to determine the optimal evolutionary model using 31 threads with the -T 31 argument. The most optimal model was WAG+R10 and was used for tree construction using the argument -m WAG+R10 with 31 threads. Branch support was calculated using Ultrafast bootstrapping with the argument -B 1000.

Tree manipulation and graphing
All processes related to tree visualization were done using Evolview72 (version 2).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript addresses an interesting and timely topic, exploring how structural modularity in fungal chitinases may drive ecological specialization between microbial antagonism and immune suppression. The work is interesting and of potential interest to the Nature Communications readership. However, the current version remains largely descriptive and lacks the depth and mechanistic insight needed to fully support some of its central claims. Several conclusions appear overstated given the evidence provided, and important experimental details, controls, and quantitative analyses are missing. Our detailed comments below highlight areas where the authors could strengthen the work through additional experiments, more rigorous data analysis, and deeper integration of their findings within the broader literature.\n- Page 4, lines 99-105. Stating that the antagonism is mediated by a GH18-CBM5 is an overstatement and should be rephrased. Although Eichfeld et al 2024 (pulished by the authors of the current manuscript) reported some evidence of a role for this GH18-CBM5 in inhibiting the pathogen B sorokiniana, many other genes were found to be induced during this interaction, therefore there is no evidence that GH18-CBM5 is THE mediator.\n- Page 6, lines 148-150: the authors should also test activity on what they later on call “protein-free Bs mycelium” (since they will show binding of the GH18-CBM5 to it, “indicating that the domain contributes to cell wall targeting”, hence the enzyme should also be active on this same substrate). A time course experiment is only provided for chitin azure, but should be done also for crab shell chitin and chitosan.\n- Fig 1: should absorbance data from assays (DNSA, chitin azure, protein binding/bradford) better be converted to more relevant values via standard curves? (also for Fig4); would be nice if TLC plate had a chito-oligo standard/ladder for comparison, not just GlcNAc\n- Proper kinetics of SiCHIT and SiCHIT2 (both with and without CBM5, and SCHIT silenced mutant) should be determined in order to properly compare the enzymes and their substrate preference.\n- Fig 1B and C, and 2 A-C, S2, Fig 4, Fig 5 need some statistics for significance (P value).\n- Page 6 and Fig 1E: the part on mass spectrometry feels a bit superficial in the main text, and the corresponding details in Methods seem to be completely missing. What mass spec instrument was used, how were samples prepared/analysed, etc? Why colloidal chitin, and not crab shell chitin or mycelium chitin? The author should also include the original mass spec results in Supp Info to prove the identity of the products, as just Fig 1E is not enough. Same for Fig 1F.\n- Conclusion in p6 line 166/167: less binding to chitosan may also be due to molecule size of chitosan compared to chitin and BS cell walls/mycelium, the authors don’t specify molecular weight (low, medium, high).\n- Fig S1: would be nice to indicate the expected molecular weight of the recombinant proteins\n- Page 7, line 165: what is this protein-free Bs mycelium and how was it made? Also, p7 line 172 “substrate binding, particular to …fungal cell wall-like chitin“ the authors just show pull down of mycelium, not specifically the chitin in the cell walls….\n- Fig 1G. This is fine, but it would be helpful to confirm it by checking bound and unbound protein using SDS-PAGE. Easy experiment to do.\n- Page 7: I do not think that treating Bs spores with Si chitinase is enough to conclude that the enzyme has an effect on pathogen growth in planta. What about the effect on Bs mycelium? The binding experiment was done on mycelium (not spores), after all, plus the mycelium may be more relevant with the actual type of in vivo interaction that takes place between the two fungi in a plant. Ideally such an experiment should be done in planta, but the alternative would be in agar plates where the chitinase is added and the growth of Bs mycelium measured. Same should be done on Si itself, to rule out negative effects of the chitinase on the producing organism.\n- There is some lack of consistency in presenting data. Fig 2 and S2 show similar results obtained from barley and Arabidopsis, respectively, yet the graphs are shown a box plots in Fig 2, and histograms in S2.\n- Page 8, line 208: please state the % of sequence identity.\n- Fig3 F) explain scale -2 to 3 L2FC; why Hv only 3 and 10 dpi?\n- What AlphaFold version was used?\n- P9 line 228 missing reference to Fig 3 FG, while no data shown for SiCHIT 3 and 4\n- Page 9, line 235-236. This statement is not in line with Fig 3I, which clearly shows that SiCHIT was induced on crab shell chitin, and SiCHIT2 was not.\n- Fig 4 B and C: a time course here is really needed, as we might be looking at plateau activity.\n- Figure 5 C: Statistical significance MUST be shown here. SiCHIT2 (without linker) should also be assessed.\n- Figure 5D is not mentioned at all in the results. Importantly, Fig 5D seems to contradict the ROS experiment. SiCHIT does not repress ROS, but does not trigger defense gene activation (over EV controls)? And SiCHIT-cbm prevents ROS generation but induces highest levels of defense gene expression? While SiCHIT2 represses ROS but induces similar levels of defense gene in barley as SiCHIT2+CBM….\n- Fig 6 not mentioned in manuscript? Or typo p11 line 302 (“Fig 5”)?\n- P11 line 281 Can you call it immune-suppressive function or is it rather the evasion of immune reaction?\n- The authors should check what happens to chitohexaose upon incubation with SiCHIT+CBM5, SiCHIT-CBM5, SiCHIT2-CBM5 and SiCHIT2+CBM5 (a time course would be best). What products are generated? Probably, the presence of CBM5 has some effect on the mechanism of action of the GH. The authors themselves state “These findings suggest that CBM5 may hinder activity on soluble substrates not by altering catalytic specificity, but by limiting access to short oligomers through spatial constraints”, however they have not provided experimental evidence. It would also be valuable to investigate the binding affinity of the enzymes for chitohexaose compared to polymeric chitin substrates. Such analyses would help elucidate the mechanistic basis for the functional differences between enzymes with or without CBM5 and their roles as effectors.\n- In addition, the authors should address how the fungal chitinase can attack the cell wall of Bipolaris while leaving the producing fungus’s own cell wall intact. This is a key, yet lacking bit of information.\n- Reference required for p3 line 66-68? P4 line 88?, (p4 line 91 own work cited), p4 line 94? P4 line 98; General: 10 out of 37 ref in total are from Zuccaro lab…", "reviewer_2": "This manuscript presents a compelling investigation into the functional divergence of GH18 chitinases in the mutualistic fungus Serendipita indica, with particular emphasis on how gain or loss of a CBM5 domain determines ecological specialization between microbial antagonism and host immune suppression. The study is conceptually interesting and addresses an important topic in fungal biology. The combination of biochemical assays, domain-swapping experiments, and host-pathogen interaction studies offers potentially valuable insights into effector modularity of symbiosis-related functions. My major concerns are outlined below:\n1. While the study demonstrates that CBM5 is required for the antifungal function of SiCHIT, the mechanistic basis of CBM5’s contribution remains superficially explored. It is unclear whether this function is unique to CBM5 or shared among other CBM family members. I suggest the authors discuss or test whether other CBM families could confer similar antifungal properties when fused to SiCHIT or SiCHIT2. This would help determine whether the observed effect is CBM5-specific or reflects a broader property of chitin-binding domains.\n2. The manuscript focuses primarily on two paralogs, SiCHIT and SiCHIT2, but it is important to understand the functional diversity of the complete GH18 chitinase repertoire in Serendipita indica. The authors mention four GH18 chitinases in the genome but provide limited functional or expression analysis beyond the two focal genes. I recommend a more systematic analysis of all GH18 family members, including their domain architecture, expression patterns under relevant conditions, and potential functional roles.\n3. A significant methodological limitation is the inconsistent inclusion of SiCHIT2 as a control across functional assays. Since SiCHIT2 is a closely related paralog lacking CBM5, it should be included alongside SiCHIT and SiCHIT-CBM5 in all comparative experiments, particularly in antifungal and immune suppression assays. Including SiCHIT2 throughout would provide a clearer baseline for interpreting how CBM5 alters function and would strengthen the conclusion regarding modularity and functional divergence.\n4. The conclusion that immune-suppressive effectors evolved from antimicrobial ancestors primarily based on the presence or absence of a CBM5 domain in a single species is an oversimplification. Domain gain or loss is only one component of effector evolution, and without a broader comparative analysis across multiple species and fungal lineages, the evolutionary trajectory remains speculative. I recommend that the authors either expand their phylogenetic and comparative domain analysis or moderate their claims regarding the evolutionary origin of chitinase modularity.", "reviewer_3": "This study, entitled “Domain gain or loss in fungal chitinases drives ecological specialization toward antagonism or immune suppression,” is an elegant study of how fungal effector proteins evolve through structural modularity and gene duplication. Focusing on two paralogous chitinases found in tandem — SiCHIT and SiCHIT2 — the authors dissect how gain or loss of the C-terminal carbohydrate-binding module CBM5 drives divergent ecological roles, including fungal antagonism and host immune suppression.\nThe fusion and deletion experiments demonstrate the modular nature of CBM5. Its presence equips chitinases for pathogen defense, while its absence tunes the enzyme for immune evasion. By tracking expression patterns, domain architecture, and protein functionality, the authors present a persuasive case for evolutionary repurposing via gene duplication and domain loss, hypothesizing a transition from an antimicrobial effector role to an effector involved in immune suppression. The work beautifully ties molecular findings to ecological specialization, showing how domain structure contributes to niche adaptation in symbiotic fungi.\nMore broadly, this study provides significant mechanistic insight into effector biology by demonstrating how structural modularity enables functional diversification. Given its potential relevance across a wide range of species, these findings have far-reaching implications. The depth and breadth of the work make it well-suited for publication in Nature Communications.\nI only have a few minor comments/suggestions\n-One of the things I was struck by was the transcriptional profile differences (regulatory rewiring) that were discovered, which are likely critical to the functional adaptation. Did the authors look at the promoter region of the paralogs to see if there was a simple explanation for this? I don’t know if this is necessary for the paper, but it would be interesting to know if the duplication included the same promoter region as part of the adaptive evolution.\n-This point may be nitpicky, but in both the methods (line 414-415) and the results (line 248), the authors mention the suppressed germination. I would argue that germination is not suppressed, but rather, the chitinase is degrading the cell wall of the new hyphae to inhibit growth rather than actually suppressing germination.\nLine 196 -Should be “…required for full SiCHIT’s antifungal…\nFigure 5A – this is difficult to follow. The authors may want to consider a different color scheme for the different treatments.\nOverall, this is an excellent story, and it was enjoyable to review.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Point-by-point response\nWe thank the Editor and all four reviewers for their careful evaluation of our manuscript and for\ntheir constructive and insightful feedback. We have addressed all comments through additional\nexperiments, expanded analyses, and substantial revision of the text. Below, we provide a\ndetailed, point-by-point response to each comment.\nResponse to reviewers\nReviewer # 1/4: This manuscript addresses an interesting and timely topic, exploring how\nstructural modularity in fungal chitinases may drive ecological specialization between\nmicrobial antagonism and immune suppression. The work is interesting and of potential\ninterest to the Nature Communications readership. However, the current version remains\nlargely descriptive and lacks the depth and mechanistic insight needed to fully support\nsome of its central claims. Several conclusions appear overstated given the evidence\nprovided, and important experimental details, controls, and quantitative analyses are\nmissing. Our detailed comments below highlight areas where the authors could strengthen\nthe work through additional experiments, more rigorous data analysis, and deeper\nintegration of their findings within the broader literature.\n1. Page 4, lines 99-105. Stating that the antagonism is mediated by a GH18-CBM5 is\nan overstatement and should be rephrased. Although Eichfeld et al 2024 (pulished\nby the authors of the current manuscript) reported some evidence of a role for this\nGH18-CBM5 in inhibiting the pathogen B. sorokiniana, many other genes were\nfound to be induced during this interaction, therefore there is no evidence that\nGH18-CBM5 is THE mediator. Reply: We agree that the previous phrasing overstated\nthe role of GH18-CBM5. We have rephrased the section to clarify that SiCHIT is not the\nsole mediator of antagonism, but rather strongly contributes to the overall antifungal\nactivity of S. indica. The revised text now states: “This antagonism involves multiple\nmolecular factors, with a GH18-CBM5 endochitinase (SiCHIT) playing a prominent\nrole by restricting Bs growth and recapitulating the protective effect of Si in planta.”\n2. Page 6, lines 148-150: the authors should also test activity on what they later on\ncall protein-free Bs mycelium (since they will show binding of the GH18-CBM5 to it,\nindicating that the domain contributes to cell wall targeting, hence the enzyme\nshould also be active on this same substrate). Reply: We agree that assessing\nenzyme activity on protein-free B. sorokiniana mycelium is important, given that SiCHIT\nbinds this substrate. We performed the additional assay and confirmed that the enzyme\nis active on protein-free fungal cell wall material. These results are now included in the\nrevised manuscript (Supplementary Fig. 14) and referenced in the Results section.\n3. A time course experiment is only provided for chitin azure, but should be done also\nfor crab shell chitin and chitosan. Reply: We performed additional time-course assays\nusing both crab shell chitin and chitosan, analogous to the chitin-azure experiment. The\nnew data are included in the revised manuscript (Fig. 1) and are described in the Results\nsection.\n4. Fig 1: should absorbance data from assays (DNSA, chitin azure, protein\nbinding/bradford) better be converted to more relevant values via standard curves?\n(also for Fig4); would be nice if TLC plate had a chito-oligo standard/ladder for\ncomparison, not just GlcNAc. Reply: We agree that converting absorbance\nmeasurements to quantitative values is preferable where possible. Accordingly, we\nrecalculated the DNSA reducing-end assays for crab shell chitin and chitosan, as well as\nthe protein–substrate binding assays, using appropriate standard curves. For the chitin-\nazure assay, such a conversion is not feasible because the substrate consists of chitin\ncovalently linked to Remazol Brilliant Violet 5R (RBV); the soluble RBV–chitin fragments\nreleased during hydrolysis do not allow the generation of a defined standard curve. In\nresponse to the request for a more informative product analysis, we removed the TLC\npanel and replaced it with a mass spectrometric characterization of the reaction products\ngenerated from crab shell chitin. This provides substantially higher resolution and more\ndetailed information than the original TLC assay.\n5. Proper kinetics of SiCHIT and SiCHIT2 (both with and without CBM5, and SCHIT\nsilenced mutant) should be determined in order to properly compare the enzymes\nand their substrate preference. Reply: We agree that kinetic parameters are important\nfor comparing substrate preferences of the different chitinases. We therefore performed\nMichaelis–Menten analyses for all relevant constructs, including SiCHIT, SiCHIT lacking\nthe CBM5, SiCHIT2, and SiCHIT2 fused to CBM5. The results are provided in the revised\nmanuscript (Supplementary Fig. 13).\nThese analyses show that the presence of the CBM5 reduces the apparent Km, consistent\nwith enhanced substrate binding, but slightly decreases Vmax, suggesting a modest\nconstraint on maximal catalytic turnover under substrate-saturating conditions. We want\nto mention here that these parameters should be interpreted cautiously because classical\nMichaelis–Menten assumptions do not fully apply to reactions involving insoluble\npolysaccharide substrates. Nevertheless, the data allow for robust comparative\nassessments and provide additional mechanistic insight into how CBM5 modulates\nenzyme performance.\n6. Fig 1B and C, and 2 A-C, S2, Fig 4, Fig 5 need some statistics for significance (P\nvalue). Reply: Each figure legend includes a dedicated section describing the statistical\ntests performed, the number of biological replicates, and the multiple-testing correction\napplied. For multi-factorial comparisons, we indicate statistically significant differences\n(adjusted p-value < 0.05) using a letter-code grouping scheme, as is standard practice.\nThis ensures consistent and transparent presentation of statistical significance across all\ndatasets.\n7. Page 6 and Fig 1E: the part on mass spectrometry feels a bit superficial in the main\ntext, and the corresponding details in Methods seem to be completely missing.\nWhat mass spec instrument was used, how were samples prepared/analysed, etc?\nWhy colloidal chitin, and not crab shell chitin or mycelium chitin? The author\nshould also include the original mass spec results in Supp Info to prove the identity\nof the products, as just Fig 1E is not enough. Same for Fig 1F. Reply: we replaced\nthe colloidal chitin sample with a mass-spectrometric product profile generated from crab\nshell chitin in Fig. 1, which represents a more relevant substrate for our study. We also\nadded the catalytically impaired SiCHITE196Q variant as a control to illustrate the expected\nresidual activity. In addition, we included a product profile for β-chitin in the Supplementary\nInformation (Supplementary Fig. 3). To address the reviewer’s request for methodological\nclarity, we substantially expanded the Methods section with full details on sample\npreparation, instrumental parameters, and data analysis. We also now provide\nrepresentative original mass spectra in Supplementary Figs. 2 and 4.\n8. Conclusion in p6 line 166/167: less binding to chitosan may also be due to molecule\nsize of chitosan compared to chitin and BS cell walls/mycelium, the authors should\nspecify molecular weight (low, medium, high). Reply: We thank the reviewer for this\nthoughtful point. The average molecular weight of the chitosan used in our assays (346\nkDa) has now been added to the Materials and Methods section. We agree that polymer\nsize and surface characteristics can influence absolute binding efficiencies. However, in\nour analysis we compare the binding behavior of the two enzymes on the same substrate\n(crab shell chitin, >75% de-acetylated chitosan, or protein-free B. sorokiniana mycelium).\nBecause each comparison is made within a single substrate type, differences in molecular\nweight do not confound our conclusions regarding the relative binding properties of the\nenzymes.\n9. Fig S1: would be nice to indicate the expected molecular weight of the recombinant\nproteins. Reply: We have added the expected molecular weights of all recombinant\nproteins to the legend of Supplementary Fig. S1 to facilitate interpretation.\n10. Page 7, line 165: what is this protein-free Bs mycelium and how was it made? Also,\np7 line 172 substrate binding, particular to fungal cell wall-like chitin the authors\njust show pull down of mycelium, not specifically the chitin in the cell walls. Reply:\nWe have now added a description of how protein-free B. sorokiniana mycelium was\ngenerated to the Materials and Methods section. We also revised the corresponding\nsection in the Results to clarify that the binding assays were performed on protein-\ndepleted fungal cell wall material enriched in chitin rather than on isolated chitin. This\nrephrasing more accurately reflects the nature of the substrate and the interpretation of\nthe binding experiments.\n11. Fig 1G. This is fine, but it would be helpful to confirm it by checking bound and\nunbound protein using SDS-PAGE. Easy experiment to do. Reply: We agree that\nconfirming the binding assay with an SDS–PAGE analysis strengthens the result. We\ntherefore analyzed bound and unbound fractions for the relevant enzyme–substrate\ncombinations by SDS–PAGE. The results, which fully support the binding patterns shown\nin Fig. 1G, are now included in the Supplementary Information (Supplementary Fig. 5).\n12. Page 7: I do not think that treating Bs spores with Si chitinase is enough to conclude\nthat the enzyme has an effect on pathogen growth in planta. What about the effect on\nBs mycelium? The binding experiment was done on mycelium (not spores), after all,\nplus the mycelium may be more relevant with the actual type of in vivo interaction that\ntakes place between the two fungi in a plant. Ideally such an experiment should be\ndone in planta, but the alternative would be in agar plates where the chitinase is added\nand the growth of Bs mycelium measured. Same should be done on Si itself, to rule\nout negative effects of the chitinase on the producing organism. Reply: We appreciate\nthe reviewer’s concern and agree that both spore- and mycelium-directed activities can\ncontribute to antifungal outcomes in planta. To clarify this point, we would like to emphasize\nthat S. indica does not interact exclusively with B. sorokiniana hyphae during colonization.\nRather, our recent imaging data show that Si hyphae establish direct physical contact with Bs\nspores on barley roots, indicating that spores represent an ecologically relevant target during\nthe early stages of the interaction. To make this explicit, we show representative confocal\nimages below that illustrate key steps of the Si–Bs interaction in planta: (i) Si hyphae coil\naround Bs spores, (ii) translocate antimicrobial effector proteins into the spores (e.g. Dld1),\nand (iii) penetrate Bs spores. These observations demonstrate that Si engages in a\nmycoparasitic attack on Bs at the spore stage, confirming that antifungal activity directed\ntoward spores is biologically meaningful and relevant for early pathogen suppression. We\nagree that a more extensive mechanistic analysis of Si-mediated antagonism, including the\nroles of additional effector proteins, would be valuable. A comprehensive dataset addressing\nthese aspects is part of a separate manuscript currently in preparation, and we therefore\nprefer not to include it here to avoid fragmenting that study. Nevertheless, we hope that the\nmicroscopy data provided below clearly show that spore-targeted antifungal activity is not an\nartificial assay choice but reflects a genuine interaction interface in planta. Finally, we note\nthat potential negative effects of SiCHIT on the producing organism have already been\nassessed in our previous work (Eichfeld et al. 2024), where treatment with the enzyme did\nnot impair S. indica growth.\n13. There is some lack of consistency in presenting data. Fig 2 and S2 show similar\nresults obtained from barley and Arabidopsis, respectively, yet the graphs are\nshown a box plots in Fig 2, and histograms in S2. Reply: We have now harmonized\nthe data presentation accordingly.\n14. Page 8, line 208: please state the % of sequence identity. Reply: We have now added\nthe corresponding percentage sequence identity to the text at the indicated location.\n15. Fig3 F) explain scale -2 to 3 L2FC; why Hv only 3 and 10 dpi? Reply: We have clarified\nboth points in the revised figure legend for Fig. 3. The legend now explains the log2 fold-\nchange (–2 to 3) scale used in panel F, and it provides the rationale for including barley\n(Hv) expression data specifically at 3 and 10 dpi, which represent the biologically most\ninformative stages for comparing SiCHIT family gene expression during barley\ncolonization.\n16. What AlphaFold version was used? Reply: We have included that information in the\nfigure legend of Figure 3.\n17. P9 line 228 missing reference to Fig 3 FG, while no data shown for SiCHIT 3 and 4.\nReply: We have now added the appropriate reference to Fig. 3F–G in the text. In addition,\nwe have included new expression data for SiCHIT3 and SiCHIT4 in the revised\nmanuscript, allowing direct comparison of all four GH18 chitinases under the relevant\nconditions.\n18. Page 9, line 235-236. This statement is not in line with Fig 3I, which clearly shows\nthat SiCHIT was induced on crab shell chitin, and SiCHIT2 was not. Reply: We thank\nthe reviewer for catching this inconsistency. We have revised the text at the indicated\nlocation to accurately reflect the expression patterns shown in Fig. 3I.\n19. Fig 4 B and C: a time course here is really needed, as we might be looking at plateau\nactivity. Reply: We performed the corresponding time-course experiments and have\nincluded the results in the Supplementary Information (Supplementary Fig. 12). The\nrelevant section of the Results has been updated accordingly to incorporate and interpret\nthese new data.\n20. Figure 5 C: Statistical significance MUST be shown here. SiCHIT2 (without linker)\nshould also be assessed. Reply: We have repeated the experiment under the same\nconditions and incorporated the new data into our analysis to strengthen the statistical\npower. Regarding the suggestion to assess SiCHIT2 without the linker region, we do not\nconsider this construct informative for our study. The native SiCHIT2 protein contains a\nshort C-terminal segment that does not connect to any additional domain and likely\nrepresents a residual linker structure retained after domain loss (see Fig. 3 and related\ntext). Because this region is part of the natural architecture of SiCHIT2, removing it would\nalso generate an artificial construct that does not reflect the endogenous enzyme. For this\nreason, and because the linker does not have a functional role comparable to the CBM5\ndomain in SiCHIT, we did not pursue a SiCHIT2 construct lacking this region.\n21. Figure 5D is not mentioned at all in the results. Importantly, Fig 5D seems to\ncontradict the ROS experiment. SiCHIT does not repress ROS, but does not trigger\ndefense gene activation (over EV controls)? And SiCHIT-cbm prevents ROS\ngeneration but induces highest levels of defense gene expression? While SiCHIT2\nrepresses ROS but induces similar levels of defense gene in barley as\nSiCHIT2+CBM. Reply: We have now incorporated explicit references to Fig. 5D in the\nResults section and substantially revised the accompanying text to clarify the\ninterpretation of these data. In response to the reviewer’s concern, we also added new\nexperimental data to Fig. 5 to strengthen the conclusions drawn from both the ROS and\ndefense gene–expression assays. The perceived discrepancy between ROS suppression\nand PR10 induction largely reflects differences in the timing and nature of the readouts.\nThe transient ROS burst occurs within minutes after chitin perception, whereas PR10\ntranscript accumulation is measured at later time points and reflects the overall interaction\noutcome rather than the immediate immune response. The slight (non-significant)\nincrease in PR10 expression observed for SiCHIT-CBM5 and SiCHIT2+link is most likely\nexplained by enhanced Si colonization at this stage, rather than by a failure to evade the\nearly immune response. In other words, the immune-evasive activity of the recombinant\nchitinases appears to provide a short-term advantage that facilitates initial colonization\n(“kick-start”), but does not persist as a long-lasting suppression of all downstream immune\noutputs in our experimental setup. We hope that the additional data and revised\nexplanation address the reviewer’s concern.\n22. Fig 6 not mentioned in manuscript? Or typo p11 line 302 (1C;Fig 5D;)? Reply:\nThank\nyou for noting this. It was a typographical error, and we have corrected it in the revised\nmanuscript.\n23. P11 line 281 Can you call it immune-suppressive function or is it rather the evasion\nof immune reaction? Reply: We agree that “immune suppression” implies a different\nmechanism. We have therefore rephrased the text.\n24. The authors should check what happens to chitohexaose upon incubation with\nSiCHIT+CBM5, SiCHIT-CBM5, SiCHIT2-CBM5 and SiCHIT2+CBM5 (a time course\nwould be best). What products are generated? Probably, the presence of CBM5 has\nsome effect on the mechanism of action of the GH. The authors themselves state.\nThese findings suggest that CBM5 may hinder activity on soluble substrates not by\naltering catalytic specificity, but by limiting access to short oligomers through\nspatial constraints, however they have not provided experimental evidence. It\nwould also be valuable to investigate the binding affinity of the enzymes for\nchitohexaose compared to polymeric chitin substrates. Such analyses would help\nelucidate the mechanistic basis for the functional differences between enzymes\nwith or without CBM5 and their roles as effectors. Reply: We have tested the release\nof reducing ends from chitohexaose at different time points and included the data in the\nsupplementary part of the manuscript (Supplementary Figure 16). Our results are in line\nwith findings on bacterial GH18 and GH19 chitinases where the CBM5 enhances\ndegradation of insoluble substrates but interferes with degradation of small, soluble\nsubstrates1,2. We have also discussed this in the current version (p 13, line 366 – 372). It\nalso aligns well with our kinetic analysis (Supplementary Figure 13) that demonstrates\nthat the CBM5 can slightly interfere with maximal reaction rates under substrate saturating\nconditions, highlighting the functional trade-off that comes with presence of the CBM5.\n25. In addition, the authors should address how the fungal chitinase can attack the cell\nwall of Bipolaris while leaving the producing fungus own cell wall intact. This is a\nkey, yet lacking bit of information. Reply: We agree that this is an important and\nfascinating question. Similar issues have been discussed for other mycoparasitic and\nantagonistic fungi3, where several mechanisms have been proposed, including cell wall\nmodifications, secretion of protective lectins, or highly localized expression of chitinases.\nA full experimental investigation of how S. indica protects its own cell wall while targeting\nB. sorokiniana would, however, constitute a substantial study on its own and is beyond\nthe scope of the current manuscript. We have now added a brief discussion of these\npossible mechanisms to the revised manuscript (p. 14, lines 394–401) to acknowledge\nthis important aspect and to highlight it as a direction for future research.\n26. Reference required for p3 line 66-68? P4 line 88?, (p4 line 91 own work cited), p4\nline 94? P4 line 98; General: 10 out of 37 ref in total are from Zuccaro lab. Reply: We\nhave added the appropriate references at all indicated positions and incorporated\nadditional citations throughout the introduction and discussion to better contextualize our\nwork within the broader literature. While some self-citation is unavoidable given our\nprevious studies on S. indica effectors and inter-microbial interactions, we have reduced\nthe overall proportion of our own references and ensured that all claims are now supported\nby external sources where available.\nReviewer #2: This manuscript presents a compelling investigation into the functional\ndivergence of GH18 chitinases in the mutualistic fungus Serendipita indica, with particular\nemphasis on how gain or loss of a CBM5 domain determines ecological specialization\nbetween microbial antagonism and host immune suppression. The study is conceptually\ninteresting and addresses an important topic in fungal biology. The combination of\nbiochemical assays, domain-swapping experiments, and host-pathogen interaction\nstudies offers potentially valuable insights into effector modularity of symbiosis-related\nfunctions. My major concerns are outlined below:\n1. While the study demonstrates that CBM5 is required for the antifungal function of\nSiCHIT, the mechanistic basis of CBM5’s contribution remains superficially explored.\nIt is unclear whether this function is unique to CBM5 or shared among other CBM\nfamily members. I suggest the authors discuss or test whether other CBM families\ncould confer similar antifungal properties when fused to SiCHIT or SiCHIT2. This\nwould help determine whether the observed effect is CBM5-specific or reflects a\nbroader property of chitin-binding domains. Reply: We thank the reviewer for this\nvaluable suggestion. We have now added a dedicated section in the Discussion (p. 12, line\n330 ff.) addressing the question of CBM5 specificity. In line with the reviewer’s point, we\nnote that while CBM5 is the canonical chitin-binding module associated with GH18\nchitinases in Basidiomycota, other CBM families with comparable chitin-binding\narchitectures or residue compositions could, in principle, substitute for CBM5 and support\nsimilar antifungal activities. The extent to which this is possible likely depends on the precise\nbinding mode and affinity contributed by each CBM type. Experimental testing of alternative\nCBMs is an interesting direction for future work and is now explicitly mentioned in the\nmanuscript.\n2. The manuscript focuses primarily on two paralogs, SiCHIT and SiCHIT2, but it is\nimportant to understand the functional diversity of the complete GH18 chitinase\nrepertoire in Serendipita indica. The authors mention four GH18 chitinases in the\ngenome but provide limited functional or expression analysis beyond the two focal\ngenes. I recommend a more systematic analysis of all GH18 family members,\nincluding their domain architecture, expression patterns under relevant conditions,\nand potential functional roles. Reply: We appreciate this suggestion and have expanded\nour analysis accordingly. We now provide additional information on all four GH18 chitinases\nin the Results section, including their domain architectures and structural predictions.\nStructural models for SiCHIT3 and SiCHIT4 have been added to the Supplementary\nInformation (Supplementary Fig. 10). Furthermore, we have integrated expression data for\nSiCHIT3 and SiCHIT4 under the relevant conditions into Fig. 3 to allow direct comparison\nwith SiCHIT and SiCHIT2. Finally, we have expanded the Discussion (p. 14, lines 377–393)\nto address the potential functional roles of each GH18 family member within the broader\ncontext of S. indica biology.\n3. A significant methodological limitation is the inconsistent inclusion of SiCHIT2 as a\ncontrol across functional assays. Since SiCHIT2 is a closely related paralog lacking\nCBM5, it should be included alongside SiCHIT and SiCHIT-CBM5 in all comparative\nexperiments, particularly in antifungal and immune suppression assays. Including\nSiCHIT2 throughout would provide a clearer baseline for interpreting how CBM5\nalters function and would strengthen the conclusion regarding modularity and\nfunctional divergence. Reply: We thank the reviewer for this comment. SiCHIT2\n(SiCHIT2+link) is included in all comparative experiments from Fig. 4 onward, including the\nbiochemical assays, antifungal assays, and immune-suppression assays. These datasets\nallow direct comparisons between SiCHIT, SiCHIT-CBM5, and SiCHIT2 variants and support\nour conclusions regarding the functional contribution of the CBM5 domain.\n4. The conclusion that immune-suppressive effectors evolved from antimicrobial\nancestors primarily based on the presence or absence of a CBM5 domain in a single\nspecies is an oversimplification. Domain gain or loss is only one component of\neffector evolution, and without a broader comparative analysis across multiple\nspecies and fungal lineages, the evolutionary trajectory remains speculative. I\nrecommend that the authors either expand their phylogenetic and comparative\ndomain analysis or moderate their claims regarding the evolutionary origin of\nchitinase modularity. Reply: Our expanded analysis shows that GH18 chitinases with and\nwithout CBM5 occur intermingled within a single fungal clade. Although our dataset does\nnot allow a definitive reconstruction of ancestral states, the observed distribution suggests\nthat, in many cases, loss of CBM5 is a more parsimonious explanation than multiple\nindependent gains within this clade, given the widespread presence of CBM5-containing\nvariants and the scattered occurrence of GH18 catalytic domains lacking CBM5. Consistent\nwith this, the outgroup of this clade consists of bacterial GH18 chitinases that carry CBM5.\nWe have adjusted the corresponding statements in the manuscript to reflect this more\ncautious interpretation and now explicitly note that the precise evolutionary history of these\nmodular arrangements remains unresolved.\nReviewer #3: This study, entitled Domain gain or loss in fungal chitinases drives ecological\nspecialization toward antagonism or immune suppression is an elegant study of how\nfungal effector proteins evolve through structural modularity and gene duplication.\nFocusing on two paralogous chitinases found in tandem SiCHIT and SiCHIT2 the authors\ndissect how gain or loss of the C-terminal carbohydrate-binding module CBM5 drives\ndivergent ecological roles, including fungal antagonism and host immune suppression.\nThe fusion and deletion experiments demonstrate the modular nature of CBM5. Its\npresence equips chitinases for pathogen defense, while its absence tunes the enzyme for\nimmune evasion. By tracking expression patterns, domain architecture, and protein\nfunctionality, the authors present a persuasive case for evolutionary repurposing via gene\nduplication and domain loss, hypothesizing a transition from an antimicrobial effector role\nto an effector involved in immune suppression. The work beautifully ties molecular findings\nto ecological specialization, showing how domain structure contributes to niche adaptation\nin symbiotic fungi.\nMore broadly, this study provides significant mechanistic insight into effector biology by\ndemonstrating how structural modularity enables functional diversification. Given its\npotential relevance across a wide range of species, these findings have far-reaching\nimplications. The depth and breadth of the work make it well-suited for publication in\nNature Communications.\n1. One of the things I was struck by was the transcriptional profile differences\n(regulatory rewiring) that were discovered, which are likely critical to the\nfunctional adaptation. Did the authors look at the promoter region of the paralogs\nto see if there was a simple explanation for this? I don’t know if this is necessary\nfor the paper, but it would be interesting to know if the duplication included the\nsame promoter region as part of the adaptive evolution. Reply: We agree that this\nis an important point. To address it, we analyzed the putative promoter regions (up to\n500 bp upstream of the start codon) of all four GH18 chitinase genes and searched for\nconserved cis-regulatory motifs. We compared these regions to the promoter\narchitecture of an expanded effector gene family in S. indica encoding DELD proteins4,\nwhich served as a reference. Whereas the upstream regions of the DELD family are\nrelatively conserved in their motif composition, our analysis revealed clear differentiation\namong the promoter regions of the four chitinase genes, consistent with the\ntranscriptional diversification observed in our expression data. We have added these\nresults to the Supplementary Information (Supplementary Fig. 11).\n2. This point may be nitpicky, but in both the methods (line 414-415) and the results\n(line 248), the authors mention the suppressed germination. I would argue that\ngermination is not suppressed, but rather, the chitinase is degrading the cell wall\nof the new hyphae to inhibit growth rather than actually suppressing germination.\nReply: We thank the reviewer for this clarification. We have rephrased the relevant\nsections in both the Methods and Results to state that the chitinase interferes with early\nhyphal outgrowth by degrading the emerging cell wall, rather than suppressing\ngermination per se.\n3. Line 196 -Should be “…required for full SiCHIT’s antifungal… Reply: We thank the\nreviewer for noting this. We have corrected the phrasing.\n4. Figure 5A – this is difficult to follow. The authors may want to consider a different\ncolor scheme for the different treatments. Reply: We agree with the reviewer that the\noriginal color scheme made the panel difficult to interpret. We have updated the colors\nfor the different treatments in Fig. 5A, and the improved contrast should make the\ndifferences much clearer.\nantifungal activities of Streptomyces griseus chitinase C. Appl Microbiol Biotechnol\n72, 1176-1184, doi:10.1007/s00253-006-0405-7 (2006).\nCBM family 5 chitin-binding domain of an alkaline chitinase from Bacillus sp. J813.\nBioscience, biotechnology, and biochemistry 76, 530-535, doi:10.1271/bbb.110835\n(2012).\nTrichoderma. Microbiology (Reading, England) 158, 26-34,\ndoi:10.1099/mic.0.052613-0 (2012).\nmetal ion homeostasis and oxidative stress. New Phytologist 227, 1174-1188,\ndoi:https://doi.org/10.1111/nph.16606 (2020).\nResponse to editor and reviewers\nWe thank the Editor and the reviewers once again for their thoughtful evaluation and\nconstructive feedback, which has substantially strengthened our manuscript. We are\npleased that the reviewers and editor agree we have adequately addressed previous\nconcerns. Below we provide point-by-point responses to the remaining comments.\nPoint-by-point response\nReviewer #1/4\nThe authors have satisfactorily addressed most of my concerns and significantly\nstrengthened the manuscript by including additional data, however there are still some\npoints that need clarification and additional evidence:\n10. I still cannot find the methods for how protein-free Bs mycelium was made. It actually\nseems that what was called “protein-free Bs mycelium” in the first manuscript has now\nbeen changed to “washed Bs mycelium” in the revised version.\nAnswer: We thank the reviewer for highlighting this point. The description of the\npreparation procedure can be found in the Methods section (lines 557–559). To avoid\nambiguity, we now consistently use the term “washed Bs mycelium”, which more\naccurately reflects the experimental procedure used.\n11. Just for clarity, Fig 1G is now 1H. Importantly, how can up to 17.5uM protein remain\nunbound, when the input only was 4uM (as stated in Supp Fig 5 and in methods)? Where\nare the input only values in Fig 1H?\nAnswer: We thank the reviewer for identifying this scaling error. We have corrected Figure\n1H scale, updated the raw data file to include input-only values, and revised the figure\naccordingly.\n24. The DNS assay is useful, however it shows that SiChit2 has only about 30% more\nactivity on chitohexaose that SiChi, yet the difference in ROS signal is massive. This must\nbe due to the type of products generated from cleavage of the substrate. The authors\nshould definitely check the profile of these products (and their abundance) and how they\nchange over time upon incubation with SiChit, SiChitE196Q, SiChit-CBM5, SiChit2+link\nand SiChit2+CBM5, using e.g. HPAEC or alternative approaches.\nAnswer: We thank the reviewer for this insightful point. We agree that the DNS assay\nprovides only an estimate of total reducing ends and therefore cannot resolve differences\nin product composition or reaction kinetics that may be critical for immune activation.\nTo address this, we performed a time-resolved analysis of chitohexaose (A6) degradation\nby all indicated chitinase variants (SiCHIT, SiCHIT^E196Q, SiCHIT–CBM5, SiCHIT2+link,\nand SiCHIT2+CBM5) using live SEC–RI–MS. This enabled monitoring of both identity and\nrelative abundance of oligomeric products (A2–A6) over time.\nAcross all active enzymes, we observed qualitatively similar product spectra, with A2 and\nA3 as the dominant end products and no evidence for distinct or unusual cleavage\nproducts. However, CBM5-lacking variants (SiCHIT–CBM5, SiCHIT2+link) depleted\nimmunogenic A6 and intermediate A4 substantially faster than CBM5-containing\ncounterparts.\nBecause chitin-triggered ROS signaling is both threshold- and time-dependent, this kinetic\nbias provides a parsimonious explanation for the observed ROS attenuation through\naccelerated depletion of immunogenic A6/A4 and accumulation of non-eliciting A2/A3 (Liu\net al., 2012; Bozsoki et al., 2017; Zang et al., 2024). We note that additional factors in\nplanta (e.g., protein stability/availability and solubility in the apoplast) could further\nmodulate the magnitude of the response.\nThese new data are provided as Supplementary Fig. 16C and are referenced in the\nrevised Results section.\nFig. 1. Product profiles of S. indica chitinases during chitohexaose (A6) degradation.\n(A) Time-resolved mass spectrometric analysis of A6 incubated with different recombinant chitinase\nvariants, monitored by live SEC-RI-MS at 37 °C. Stacked bars represent relative MS signal intensities of\noligomeric products (A2–A6) over time. (B) Average number of cleavage events per A6 molecule calculated\nfrom product distributions shown in (A). (C) Relative enzymatic activity of SiCHIT and SiCHIT2 variants with\nor without CBM5, normalized to the corresponding full-length enzyme. Percentages were calculated from\nthe data shown in (B).\nReviewer #2\nMy questions have been largely addressed. However, I would like to raise one remaining\nconcern regarding the interpretation of the CBM5-associated phenotype. The current\nconclusion appears overstated. From a biochemical perspective, the primary function of\nthe CBM5 domain is to enhance substrate binding and facilitate the proximity of the\ncatalytic GH18 domain to chitin. While the resulting phenotypic consequences may\nmanifest as immune suppression or microbial antagonism, these effects are likely\nsecondary outcomes of improved substrate engagement rather than evidence that CBM5\nhas evolved a fundamentally new immune-related function. Therefore, I suggest the\nauthors moderate their claims about functional “gain” or “loss”, in evolution and clarify that\nthe core activity remains chitin hydrolysis, with CBM5 mainly modulating efficiency and\ncontext of substrate targeting.\nAnswer: We thank the reviewer for this thoughtful comment and agree that CBM5\nprimarily functions as a carbohydrate-binding module that enhances substrate\nengagement for the GH18 catalytic domain, rather than conferring a new catalytic activity.\nAt the same time, our data show that the presence of CBM5 strongly enhances antifungal\nactivity and plant protection by SiCHIT and is sufficient to confer these antagonistic\nfunctions when fused to the paralog SiCHIT2. Conversely, CBM5-lacking variants more\nefficiently deplete soluble immunogenic chitooligomers, suppress chitin-triggered ROS,\nand promote early root colonization. Thus, CBM5 does not change product specificity but\nshifts substrate targeting and reaction dynamics in a context-dependent manner, leading\nto distinct outcomes in microbe–microbe and host–microbe interactions. We have\nmoderated our wording throughout the manuscript, including the title, and explicitly\nemphasize that the core activity remains chitin hydrolysis. We added the following\nclarification to the Discussion: “Biochemically, our data support the view that CBM5\nprimarily enhances substrate engagement and positioning of the GH18 catalytic domain\non crystalline chitin. Its presence or absence is sufficient to shift the efficiency and context\nof chitin hydrolysis in ways that translate into distinct biological outcomes, ranging from\nmicrobial antagonism to stronger attenuation of chitin-triggered host immunity”\nReviewer #3\nThe authors have adequately addressed the reviewers concerns.\nAnswer: We are happy that the current version of the manuscript meets your\nexpectations. Thank you again for your helpful comments and feedback that helped to\nimprove our study.\nReferences\nZ. Bozsoki, J. Cheng, F. Feng, K. Gysel, M. Vinther, K.R. Andersen, G. Oldroyd, M.\nBlaise, S. Radutoiu, & J. Stougaard, Receptor-mediated chitin perception in legume\nroots is functionally separable from Nod factor perception, Proc. Natl. Acad. Sci. U.S.A.\n114 (38) E8118-E8127, https://doi.org/10.1073/pnas.1706795114 (2017)\nTingting Liu et al. Chitin-Induced Dimerization Activates a Plant Immune Receptor.\nScience336,1160-1164(2012). DOI:10.1126/science.1218867\nZhang J, Sun J, Chiu CH, Landry D, Li K, Wen J, Mysore KS, Fort S, Lefebvre B, Oldroyd\nGED, Feng F. A receptor required for chitin perception facilitates arbuscular mycorrhizal\nassociations and distinguishes root symbiosis from immunity. Curr Biol. 2024 Apr\n22;34(8):1705-1717.e6. doi: 10.1016/j.cub.2024.03.015. Epub 2024 Apr 3. PMID:\n38574729; PMCID: PMC11037463."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have satisfactorily addressed most of my concerns and significantly strengthened the manuscript by including additional data, however there are still some points that need clarification and additional evidence:\n- Point 10. I still cannot find the methods for how protein-free Bs mycelium was made. It actually seems that what was called “protein-free Bs mycelium” in the first manuscript has now been changed to “washed Bs mycelium” in the revised version.\n- Point 11. Just for clarity, Fig 1G is now 1H. Importantly, how can up to 17.5uM protein remain unbound, when the input only was 4uM (as stated in Supp Fig 5 and in methods)? Where are the input only values in Fig 1H?\n- Point 24. The DNS assay is useful, however it shows that SiChit2 has only about 30% more activity on chitohexaose that SiChi, yet the difference in ROS signal is massive. This must be due to the type of products generated from cleavage of the substrate. The authors should definitely check the profile of these products (and their abundance) and how they change over time upon incubation with SiChit, SiChitE196Q, SiChit-CBM5, SiChit2+link and SiChit2+CBM5, using e.g. HPAEC or alternative approaches.", "reviewer_2": "Thank you for the authors’ careful and constructive responses.", "reviewer_3": "The authors have adequately addressed the reviewers concerns.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Answer: We thank the reviewer for highlighting this point. The description of the preparation procedure can be found in the Methods section (lines 557–559). To avoid ambiguity, we now consistently use the term “washed Bs mycelium”, which more accurately reflects the experimental procedure used.\nAnswer: We thank the reviewer for identifying this scaling error. We have corrected Figure 1H scale, updated the raw data file to include input-only values, and revised the figure accordingly.\nAnswer: We thank the reviewer for this insightful point. We agree that the DNS assay provides only an estimate of total reducing ends and therefore cannot resolve differences in product composition or reaction kinetics that may be critical for immune activation. To address this, we performed a time-resolved analysis of chitohexaose (A6) degradation by all indicated chitinase variants (SiCHIT, SiCHIT^E196Q, SiCHIT–CBM5, SiCHIT2+link, and SiCHIT2+CBM5) using live SEC–RI–MS. This enabled monitoring of both identity and relative abundance of oligomeric products (A2–A6) over time. Across all active enzymes, we observed qualitatively similar product spectra, with A2 and A3 as the dominant end products and no evidence for distinct or unusual cleavage products. However, CBM5-lacking variants (SiCHIT–CBM5, SiCHIT2+link) depleted immunogenic A6 and intermediate A4 substantially faster than CBM5-containing counterparts. Because chitin-triggered ROS signaling is both threshold- and time-dependent, this kinetic bias provides a parsimonious explanation for the observed ROS attenuation through accelerated depletion of immunogenic A6/A4 and accumulation of non-eliciting A2/A3 (Liu et al., 2012; Bozsoki et al., 2017; Zang et al., 2024). We note that additional factors in planta (e.g., protein stability/availability and solubility in the apoplast) could further modulate the magnitude of the response. These new data are provided as Supplementary Fig. 16C and are referenced in the revised Results section.\nAnswer: We thank the reviewer for this thoughtful comment and agree that CBM5 primarily functions as a carbohydrate-binding module that enhances substrate engagement for the GH18 catalytic domain, rather than conferring a new catalytic activity. At the same time, our data show that the presence of CBM5 strongly enhances antifungal activity and plant protection by SiCHIT and is sufficient to confer these antagonistic functions when fused to the paralog SiCHIT2. Conversely, CBM5-lacking variants more efficiently deplete soluble immunogenic chitooligomers, suppress chitin-triggered ROS, and promote early root colonization. Thus, CBM5 does not change product specificity but shifts substrate targeting and reaction dynamics in a context-dependent manner, leading to distinct outcomes in microbe–microbe and host–microbe interactions. We have moderated our wording throughout the manuscript, including the title, and explicitly emphasize that the core activity remains chitin hydrolysis. We added the following clarification to the Discussion: “Biochemically, our data support the view that CBM5 primarily enhances substrate engagement and positioning of the GH18 catalytic domain on crystalline chitin. Its presence or absence is sufficient to shift the efficiency and context of chitin hydrolysis in ways that translate into distinct biological outcomes, ranging from microbial antagonism to stronger attenuation of chitin-triggered host immunity”\nAnswer: We are happy that the current version of the manuscript meets your expectations. Thank you again for your helpful comments and feedback that helped to improve our study."}]

Decision letter