Siderophore-producing Bacillus and free-living nematodes are associated with soil suppressiveness to banana root-knot nematodes

Domain: neuroscience
Published: 2026-02-12
Editorial outcome: major_revision

Abstract
The control of soil-borne diseases is crucial for ensuring global food security. Here, we investigate the impact of the root-knot nematode (Meloidogyne) on banana continuous cropping over a period of 11 years. The results show significant root infestation initially, but disease incidence declined markedly from the 7th cropping year onwards. Soil community profiling revealed that this intriguing onset of nematode suppressiveness was associated with changes in free-living nematode populations and rhizosphere microbiome composition. Rhizosphere microbiome analyses and strain isolation pinpointed Bacillus velezensis as a keystone taxon in soil suppressiveness to Meloidogyne. Genomics, metabolomics and bioassays validated the suppressive effects of B. velezensis against Meloidogyne and identified the siderophore bacillibactin as key metabolite with repellent and nematicidal activities. By integrating long-term field studies with multi-omics approaches, this study uncovered co-occurring increases in specific rhizobacterial genera and free-living nematodes associated with reduced root-parasitic nematode populations, offering valuable insights for sustainable agriculture.

Introduction
Soil-borne diseases pose a persistent and escalating threat to global food security, particularly under intensive agricultural systems1,2. Among soil-borne pathogens, plant parasitic nematodes (PPNs) are especially destructive, accounting for an estimated 10% loss in global crop yields annually1,3,4. This problem is exacerbated by the widespread practice of continuous cropping where the repeated cultivation of a single crop leads to the buildup of soil pathogens, including PPNs5–7. For example, continuous monoculture of banana (Musa) frequently results in the proliferation of root-knot nematodes, notably those from the genus Meloidogyne, which cause substantial yield losses8. Current management practices rely heavily on chemical nematicides8–10. However, their environmental and ecological consequences have spurred urgent calls for more sustainable alternatives.

One promising avenue is the exploitation of natural disease-suppressive soils, where native soil microbiota suppresses pathogens without the need for external inputs. These suppressive effects often emerge over time during continuous cropping, particularly following early disease outbreaks6,11. Suppressive soils are typically characterized by the enrichment of specific microbial taxa in the rhizosphere that interfere with key stages in the pathogen’s lifecycle12–14. For example, genera such as Pseudomonas, Purpureocillium and Pochonia have been implicated in the biological suppression of soybean cyst nematodes6. Several root-knot nematode-suppressive soils have been found in nature, most studies are not focused on those formed under long-term nematode infestation in continuous cropping conditions15. Numerous microbial strains isolated from nematode-suppressive soils and have been well investigated and commercialized16,17. However, despite the identification of some antagonistic microbial genera18–20, the mechanistic understanding of microbiome-mediated suppression of Meloidogyne under long-term monoculture remains largely elusive. In particular, the multipartite interactions between the rhizosphere microbiome, other soil microbiota and Meloidogyne have received little attention.

To address this knowledge gap, we monitored a long-term banana monoculture system as a model to uncover the (micro)biological mechanisms underlying natural suppression of Meloidogyne-induced root-knot disease. To unravel the key players associated with induced suppressiveness to root-knot disease, we focused our analyses on the putative roles of free-living nematodes (FLNs, including bacterivores, fungivores, and omnivore-predators) and the banana rhizosphere microbiome. FLNs can enhance plant performance by increasing carbon and nutrient cycling and suppressing plant pathogens, making most FLNs beneficial to plants21,22. FLNs are also considered non-target organisms in PPN control, with Caenorhabditis elegans being a prominent member that is widely used as a biological model for soil toxicity studies23. Considering the crucial role of the microbiome in plant fitness and disease control, we hypothesize that prolonged continuous cropping reshapes both the nematode community and the rhizosphere microbiome, leading to increased abundance of FLNs and specific microbial taxa with complementary suppressive activities.

To test our hypothesis, we examined a long-term (11 years) field trial, tracking the incidence of root-knot nematode disease over time (Fig. 1). We combined phenotypic assessments of disease incidence and high-throughput sequencing to analyze changes in soil nematode community and banana rhizosphere microbiome. Our results revealed a significant decline in disease incidence after the 7th year, coinciding with shifts in FLNs diversity and enrichment of Bacillus species in the rhizosphere. Further isolation and functional validation demonstrated that Bacillus velezensis was a keystone taxon in disease suppression, with the siderophore bacillibactin identified as a key metabolite exhibiting both repellent and nematicidal effects on Meloidogyne. Importantly, bacillibactin showed minimal toxicity to C. elegans, underscoring its potential as a selective biocontrol agent. Together, our findings highlight evidence of root-knot nematode suppression through the synergistic effect of free-living nematodes and siderophore-secreting rhizobacteria. Our study advances the understanding of microbiome-mediated soil suppressiveness and offer eco-friendly strategies for sustainable nematode management in intensive cropping systems.

Results
Induction of suppressiveness to root-knot nematode disease during continuous banana cropping
To determine the effect of different cropping times on banana root-knot nematode disease occurrence, we collected the samples from four groups of banana plantations with continuous cropping for 1 (Y1), 4 (Y4), 7 (Y7), and 10 years (Y10) in 2016; 2 (Y2), 5 (Y5), 8 (Y8), and 11 (Y11) years in 2017, and investigated root-knot nematode disease incidence (Fig. 1a and Fig. 2a). Root-knot nematode disease incidence showed a parabolic pattern, peaking at 90–96% in plantations of 2, 4, and 5 years of age (Y2, Y4, Y5), followed by a significant decline to 40–50% after 7, 8, 10 and 11 years (Y7, Y8, Y10, Y11). A similar trend was observed for gall index results (Fig. 2b, c). Although different sets of cropping years were sampled in 2016 and 2017, combining the data showed consistent patterns in disease incidence across years, supporting the robustness of the space-for-time substitution design. Total soil nematode abundance displayed a concomitant increase during continuous banana cultivation, with the lowest levels observed in Y1 and Y2 (approximately 1800 individuals per 100 g dry soil), while in Y11 the highest nematode abundances existed, approximately 4 times higher than in Y1 and Y2 (Fig. 2d). Collectively, these results show outbreaks of root-knot nematode disease of banana in the first 5 years of continuous cropping, followed by a decline in disease incidence and gall index from year 7 onwards.

Association of free-living soil nematode abundance and root-knot suppressiveness
To investigate the potential involvement of soil nematode abundance and community structure in induced suppressiveness to root-knot disease of banana caused by Meloidogyne, we integrated morphological assessments (in 2016, including Y1, Y4, Y7, and Y10) and high-throughput sequencing (in 2017, including Y2, Y5, Y8, and Y11) (Fig. 1a). For the results in 2016, we observed that the Shannon index was lower in Y1 compared to Y4, Y7, and Y10, indicating a continuous cropping effect on nematode α diversity (Fig. 3a). Unconstrained principal coordinate analysis (PCoA) showed that long-term continuous banana cropping significantly influenced soil nematode communities in 2016 (Adonis; R2 = 0.41; p < 0.001) and in 2017 (Adonis; R2 = 0.38; p < 0.001), with a distinct separation between short-term and long-term cropping (Fig. 3b and Supplementary Fig. 1a).

Analysis of nematode trophic groups in 2016 revealed that plant-parasites (Pp) increased in relative abundance from 13.9% in year 1 (Y1) to 50.5% in year 4 (Y4), then decreased to 37.5% in year 7 (Y7) and 30.9% in year 10 (Y10) (Fig. 3c). Among the FLNs (putatively beneficial nematodes), bacterivores (Ba) were most abundant in Y1 at 73.6% while decreasing to 30–40% in subsequent cropping times; the relative abundance of fungivores (Fu) increased with continuous cropping, rising from 8.4% in Y1 to nearly 20% in Y7 and Y10, omnivores-predators (OP) showed similar changes, increasing from 3.8% in Y1 to nearly 11% in Y7 and Y11 (Fig. 3c).

The relative abundance of Meloidogyne showed a similar trend with Pp, increasing from 2% in Y1 to 6.5% in Y4 and decreasing to 1.3% in Y10 (Fig. 3d). Under the 18S rRNA amplicon sequencing results, we also found that Meloidogyne was most abundant in Y2 and Y5 at 8.0-13.0%, while decreasing to 0.1-1% in Y8 and Y11 (Supplementary Fig. 1b). Combining the nematode trophic groups with Meloidogyne relative abundance, a significant positive association was found between the Meloidogyne relative abundance and Pp relative abundance (Pearson’s r = 0.63, p < 0.001), whereas negative associations were observed between Meloidogyne relative abundance and Fu relative abundance (Pearson’s r = -0.52, p = 0.0029) and OP relative abundance (Pearson’s r = -0.47, p = 0.0083). No significant association was found between Meloidogyne relative abundance and Ba relative abundance (Pearson’s r = -0.2, p = 0.29) (Fig. 3e). Collectively, these results indicated that long-term continuous banana cropping is associated with a compositional shift in the soil nematode community, with a reduced abundance of PPNs (especially Meloidogyne) and an increase in abundance of FLNs (especially Fu and OP).

Long-term continuous cropping enriches specific rhizobacterial genera with Bacillus as a key taxon
To explore the putative effects of the banana root microbiome on root-knot nematode disease during continuous banana cropping, we investigated the changes in abundance and community structure of rhizosphere bacteria (Fig. 1b). Results of 16S rRNA gene amplicon sequencing showed that the bacterial community composition of the banana rhizosphere changed during continuous banana cropping with significantly higher diversity in Y5, Y8, and Y11 (7.9–8.0 of Shannon index and 4324–4382 of Chao1) compared to Y2 (7.7 of Shannon index and 3581 of Chao1) (Fig. 4a). Consistent changes were observed in bacterial copy numbers, which increased with continuous cropping, reaching the highest abundance of 13.6 × 108 copies per gram of soil in Y11 (Fig. 4b). Moreover, PCoA based on Bray-Curtis distances revealed changes in bacterial community structure with prolonged cropping (Adonis; R2 = 0.32; p < 0.001) (Fig. 4c).

Co-occurrence network analyses of 16S amplicon sequence variants (ASVs) detected in the early (Y2, Y5) and late (Y8, Y11) stages of cropping revealed two modules, each associated with relatively high proportions of indicator ASVs, i.e. bacterial taxa that responded most significantly to cropping time (Spearman’s ρ > 0.6, p < 0.01) (Fig. 4d). Notably, ASVs assigned to Y2, Y5 were predominantly located in module 1 (77 ASVs) and distinct from module 2 (117 ASVs), which primarily contained ASVs assigned to Y8 and Y11 (Fig. 4e and Supplementary Data 1). Each module consisted of multiple phyla/genera: Actinobacteriota was more abundant in module 1, whereas Acidobacteriota and Proteobacteria (e.g. genus Allorhizobium-Neorhizobium-Pararhizobium-Rhizobium; Mesorhizobium) were more abundant in module 2 (Fig. 4f and Supplementary Table 1). Four phyla/genera, including Firmicutes (e.g. genus Bacillus), Gemmatimonadota, Myxococcota, and Nitrospirota (e.g. genus Nitrospira) exclusively appeared in module 2, whereas Planctomycetota was observed only in module 1 (Fig. 4f and Supplementary Table 1). Positive associations were observed between module 1 and Meloidogyne relative abundance (Pearson’s r = 0.4; p = 0.023), whereas significant negative associations were found between module 2 and Meloidogyne relative abundance (Pearson’s r = -0.37; p = 0.037) (Fig. 4g).

To identify bacterial taxa potentially involved in root-knot nematode disease suppressiveness, we selected the top 10 ASVs that responded most significantly to the cropping time and were most enriched in Y8 and Y11 as compared to Y2 and Y5. Three specific ASVs (ASV6738, ASV1467, ASV7704) showed significantly negative correlations with Meloidogyne relative abundance and disease incidence. Notably, ASV1467 had the strongest negative association (Spearman’s ρ = -0.64) with Meloidogyne relative abundance (Fig. 5a and Supplementary Fig. 2). Independently, we conducted random forest modelling with the indicator ASVs to further predict the potential link between the main ASV predictors of Meloidogyne relative abundance. Consistently, the results indicated that the most important predictor of Meloidogyne was ASV1467 (9.5%) (Supplementary Fig. 3a). ASV1467 was taxonomically delineated as Bacillus and enriched in the banana rhizosphere in Y8 (0.24%) and Y11 (0.27%) as compared to Y2 (0.15%) and Y5 (0.07%) (Fig. 5b). These findings suggest a potential role of ASV1467 in root-knot nematode suppressiveness.

Bacillus velezensis and its siderophore bacillibactin suppress M. incognita in banana via repellent and nematicidal activities
To test the protective role of ASV1467 against M. incognita in banana, we isolated a total of 190 rhizobacteria. Based on 16S rRNA sequencing of these isolates, strain Y11.1, isolated from banana rhizosphere samples of Y11, had an identical 16S rRNA sequence (451 bp) as ASV1467 (Supplementary Fig. 3b). We then sequenced the genome of Bacillus sp. Y11.1 and whole-genome phylogenetic analyses revealed that strain Y11.1 belongs to the B. velezensis group (Fig. 5c and Supplementary Fig. 4). Subsequent in vitro assays showed that B. velezensis Y11.1 exhibited lethal effect on M. incognita, with relative mortality rates reaching 69.72% at 48 h (Supplementary Fig. 5).

To experimentally validate that B. velezensis Y11.1 can suppress root-knot nematodes in soil, we conducted two greenhouse experiments (Fig. 1b and Fig. 5d). When banana plants were cultivated in sterilized soil, a treatment with cell suspensions of B. velezensis Y11.1 promoted banana growth and significantly decrease in M. incognita infection and reproduction (Fig. 5e and Supplementary Fig. 6a). Gall and egg mass numbers were reduced by 46.2% and 69.6%, respectively. Similar outcomes were found under natural field soil conditions, with B. velezensis Y11.1 reducing galls numbers by 59.4%, while enhancing abundance of total soil nematodes by 45.6% and promoting banana growth (Fig. 5f and Supplementary Fig. 6b). Quantitative PCR analysis confirmed a significant reduction in Meloidogyne density in soil treated with B. velezensis Y11.1 by 48.7%. While B. velezensis Y11.1 application to natural soil significantly reduced RKN density, it concurrently increased total nematode abundance due to the proliferation of non-parasitic trophic groups. Thus, the apparent increase in total nematodes does not reflect an increase in RKN, but rather the selective suppression of the pathogen alongside enrichment of free-living nematodes. Our results affirm the suppressive effect of B. velezensis Y11.1 against M. incognita in banana plants and its ability to promote banana growth.

To further identify which metabolites from B. velezensis Y11.1 inhibit M. incognita, genome mining revealed several secondary metabolites biosynthetic gene clusters (BGCs), including 13 BGCs associated with the biosynthesis of non-ribosomal peptides, polyketides, terpenes, and post-translationally modified peptides (RiPPS). Based on earlier discoveries highlighting potent nematode-suppressive attributes of siderophores24,25, we focused on the siderophores produced by B. velezensis Y11.1 (Fig. 1c). We experimentally showed that Y11.1 supernatants from iron-limited culture conditions (containing siderophore(s)) caused 21.6% more mortality than supernatants from iron-rich culture conditions (without siderophore) (Fig. 6a), suggesting lethal effects of Y11.1 siderophore(s) on Meloidogyne juveniles. Next, we directed our attention specifically to bacillibactin, a catecholate-type siderophore predicted for one of the BGCs identified in the Y11.1 genome (Supplementary Fig. 7a). The supernatant of Y11.1 cultivated in iron-limited SSM broth, contained 54.7 μM bacillibactin (Supplementary Fig. 7b, c). We next examined the inhibitory effects of bacillibactin on Meloidogyne. We found that the purified bacillibactin at 500 μM exhibited lethal effects on Meloidogyne juveniles, resulting in relative mortality rates of 11.51% at 48 hrs. In contrast, we found no significant difference in C. elegans mortality at various concentrations of bacillibactin compared to the controls (Fig. 6b). We also observed that the Y11.1 supernatant from iron-limited SSM and the purified bacillibactin repelled Meloidogyne juveniles: highest chemotaxis indices were found at -0.83 of SSM and -0.30 at 500 μM bacillibactin, respectively (Fig. 6c and Supplementary Fig. 8). In contrast, at concentrations of 50–500 μM, bacillibactin attracted C. elegans with an increase in the chemotaxis index from 0.06 to 0.61 (Fig. 6c). To experimentally validate that bacillibactin plays an important role in inhibiting Meloidogyne infection of banana roots, we conducted two greenhouse experiments (Fig. 6d). We found that both in sterilized soil and natural soil conditions, adding 40 μmol/kg bacillibactin every 14 days significantly reduced the gall, egg mass numbers, and the nematode abundance in the root, and also promoted banana growth (Fig. 6e, f and Supplementary Fig. 9). These results indicate that the siderophore bacillibactin of B. velezensis Y11.1 plays a role in alleviating root-knot nematode infection and reproduction by repelling and causing mortality in Meloidogyne juveniles, but attracts the free-living nematode C. elegans without causing lethality.

Methods
Description of experimental site and sampling procedure
The study was conducted in a banana cultivation field located in Nanning, Guangxi, China (23°05´N, 107°82´E), characterized by a subtropical climate with an average annual temperature of 21.8 °C and mean annual precipitation of 1301 mm. The soil type in this area is a highly weathered Udic Ferralsol.

The field experiment spanned from March 2016 to October 2017, focusing on banana plants of the Musa AAA Cavendish cv. Williams B6 variety (Fig. 1a). Cultivation practice, including fertilization, and agronomic management remained consistent across the different time series plantations. Samples were collected using a space-for-time substitution approach. In 2016, we sampled four groups of banana plantations that had been continuously cropped for 1 (Y1), 4 (Y4), 7 (Y7), and 10 (Y10) years. In 2017, we re-sampled these same four groups of plantations, which by then had reached 2 (Y2), 5 (Y5), 8 (Y8), and 11 (Y11) years of continuous cropping (Fig. 1a). This design was chosen due to logistical constraints of banana constant growth and large in size that prevented sampling every cropping duration in both years, but it allowed us to represent the full temporal sequence of continuous cropping across sites. For each cropping duration (e.g., 1, 4, 7, or 10 years), 10 plantations (ranging from 1 to 2 ha) were randomly selected, and composite samples were obtained from five uniformly growing banana plants (200 g soil sample and 50 g root sample) using a five-point method. These 10 samples were treated as biological replicates for statistical analysis within each site. Sampling was conducted at three time points each year (March, July, and October) to capture seasonal variation in nematode infectivity and rhizosphere dynamics. Each field in 2016 was re-sampled in 2017 (e.g., Y1/Y2, Y4/Y5), ensuring consistency in spatial location across years. Initial analyses revealed no consistent differences across seasons or years, and therefore disease indices were averaged across seasons for each cropping year to reduce complexity and improve clarity in data presentation. Due to logistical constraints, we were unable to sample multiple independent plantations per cropping duration. Thus, this design uses a space-for-time substitution approach and reflects within-site variation.

The root samples were collected per banana plant (50 g) for gall assessment. Galling of root systems was rated using a 0–5 rating scale43. The gall rate was assigned as: 0 = 0%, 1 = 1–15%, 2 = 15–30%, 3 = 30–50%, 4 = 50–75%, and 5 = 75–100% root galled, gall index = (gall rate × plant numbers)/(the highest gall rate × all plant numbers) × 100. For the Y1 field, where banana seedlings were planted in August 2016, the first sampling took place in October. Upon collection, soil samples were mixed thoroughly and sieved through a 2-mm sieve. The rhizosphere soil samples were stored at -80 °C for further analysis.

Analysis of soil nematode diversity
The soil nematodes were extracted from 100 g of soil using the modified Baermann funnel method and counted under a stereomicroscope44. After counting the soil nematode abundance, we identified nematodes (in 2016) further assigned into genus and four trophic groups, as bacterivores (Ba), fungivores (Fu), herbivores (He; Plant-parasitic, Pp), omnivores-predators (OP)45. A total of 60 nematode genera were identified, including 20 Ba, 13 Pp, 6 Fu, and 21 OP (Supplementary Tables 2-4). All of the soil nematodes were determined as colonizer–persister groups (cp1-5)46. Alpha diversity of the nematode community was determined by the Shannon-Wiener diversity index (Shannon index), which was calculated as following: Shannon index = -∑Pi × ln Pi, in which ‘Pi’ represents the proportion of individuals of the ‘i’ group in the community.

In 2016 (Y1, Y4, Y7, Y10), nematode communities were assessed by morphological identification, which provides accurate relative abundances but is limited in throughput. In 2017 (Y2, Y5, Y8, Y11), high-throughput sequencing was applied to analyze nematode communities and, in parallel, rhizosphere microbial communities from the same soils. This sequential design was chosen to build on the first year’s observations and to enable integrated microbial–nematode analyses, while reflecting practical constraints on resources and field logistics.

Soil DNA extraction, 16S rRNA gene and 18S rRNA gene amplicon sequencing
The 16S rRNA gene and 18S rRNA gene amplicon sequencing were performed to examine the impact of continuous cropping on rhizosphere bacterial and nematode compositions, respectively. Total DNA was isolated from 500 mg rhizosphere soil sample using the FastDNA® Spin Kit for Soil (MP Biomedicals, Solon, USA) according to the manufacturer’s instructions. The hypervariable region V3-V4 of the bacterial 16S rRNA gene was amplified with primers 338F (5’-ACTCCTACGGGAGGCAGCAG-3’) and 806R (5’-GGACTACHVGGGTWTCTAAT-3’) by an ABI GeneAmp® 9700 PCR thermocycler (Applied Biosystems, Foster City, USA). Illumina MiSeq PE300 platform (Illumina, San Diego, USA) was used to sequence by the Majorbio Bio-Pharm Technology Co. Ltd. (Shanghai, China). For 18S rRNA gene sequencing of soil nematodes, primers NF1F (5′-GGTGGTGCATGGCCGTTCTTAGTT-3′)47 and NR (5′AGCGACGGGCGGTGTGTACAAA-3′)48 were used. 18S rRNA primers were selected to specifically target soil nematodes, and therefore fungal taxa were underrepresented in the amplicon dataset. The amplification and sequencing were performed as described above.

The preprocessing of amplicon sequence data was conducted using usearch 11.0.667. Following the removal of PCR primers and low-quality sequences, the bacterial community samples yielded 25519-40718 reads per group, while the soil eukaryote community samples ranged from 17852 to 48698 reads (275-19389 reads were taxonomically assigned to nematozoa per group). The processed reads were denoised and merged to generate amplicon sequence variants (ASVs). ASVs were taxonomically classified using the SINTAX algorithm in vsearch-2.23.0 and the SILVA release 138 database, with the exclusion of mitochondrial and chloroplast reads. Additionally, features with a total abundance of less than three across all samples were filtered out. This process resulted in 8725 bacterial features (Supplementary Data 2) derived from the 16S rRNA V3-V4 region and 14 nematode features (Supplementary Data 3) from the 18S rRNA V4 region. 16.5% of the 18S rRNA reads were identified to 29 nematode taxa (Supplementary Data 4 and Supplementary Table 5).

Quantitative PCR of rhizobacteria
Total bacterial abundance in the rhizosphere was quantified using quantitative PCR (qPCR) with primers 338F/806R for the assessment of total soil bacterial density. The total soil DNA was extracted as described above, and subsequent to DNA quantification and concentration assessment, the qPCR assays were performed according to the standard protocols by Majorbio Bio-Pharm Technology Co. Ltd. (Shanghai, China). Briefly, the PCR reaction mixture (20 μL) contained 10 μL ChamQ SYBR Color qPCR Master Mix (2X), 0.4 μL each primer (5 μM), 2 μL template DNA (10 ng/μL) and 7.2 μL ddH2O, with each DNA sample analyzed in three replicates. Following confirmation of the amplified fragment’s specificity by melt analysis and agarose gel electrophoresis. The thermal cycling profile include a first step at 95 °C for 5 min, followed by 40 cycles of 95 °C for 30 s and 56 °C for 30 s, with a final extension at 72 °C for 40 s, and extension at 72 °C for 10 min. The results were expressed as copies/g soil by calculating the copy number of the target fragment from the standard.

Isolation and identification of rhizobacteria
Rhizobacteria were isolated from the rhizosphere of Y8 and Y11, where the disease index decreased (Fig. 1b). Briefly, one gram of the soil samples was agitated in 9 mL sterilized deionized water for 10 min. The resulting microbial suspension was serially diluted tenfold and spread onto Luria-Bertani (LB) agar plates. Incubation was carried out at 30 °C for 48 h in darkness. Colonies with distinct morphologies were selected and re-streaked on LB plates for further purification. The final collection consisted of 190 rhizobacteria isolates. Purified isolates were cultured in liquid LB medium at 30 °C with shaking at 180 r.p.m. The 16S rRNA gene of the rhizobacteria isolates was amplified using 27F (5’-AGAGTTTGATCMTGGCTCAG-3’) and 1492R (5′-TACGGYTACCTTGTTACGACTT-3′) to generate phylogenetic information. The crude electropherogram was analyzed by the MEGA v.11.0 software. The isolated strains sequences were compared with other sequences using the BLAST in the NCBI database followed by phylogenetic analysis with the neighbor-joining (NJ) method in the MEGA software. Y11.1 sequence was deposited at GenBank, and the accession numbers was PP789636.

Total genomic DNA extraction and identification of Bacillus Y11.1
The genome of Bacillus Y11.1 isolate was sequenced using a combination of PacBio RSII Single Molecule Real Time (SMRT) and Illumina sequencing platforms. Bacillus Y11.1 and 19 other strains with complete genome sequences were selected to construct a phylogenetic tree with 30 housekeeping genes as phylogenetic markers, and employed the NJ method with MEGA v.11.0. All analyses were performed on the I-Sanger Cloud Platform from Shanghai Majorbio. The genome mining for biosynthetic gene clusters of antimicrobial compounds was conducted with antiSMASH 7.049. Siderophore biosynthesis was further investigated with detailed blast and literature comparisons.

Assessing the nematicidal activity of B. velezensis Y11.1
To assess the nematicidal activity of B. velezensis Y11.1, we conducted in vitro experiments using LB medium. Cultures were grown at 30 °C with shaking at 180 r.p.m until reaching OD600 = 1.0. After filtering through a 0.22 μm membrane, the culture was diluted with LB medium to concentrations of 1x, 0.5x, and 0.1x. All solutions were supplemented with 100 μg/mL rifampicin, 20 μg/mL penicillin, and 50 μg/mL streptomycin to avoid contamination. A total of approximately 100 M. incognita J2s in a 10 μL suspension and 500 μL of different dilutions of LB culture medium were added into 24-well culture plates, with distilled water (CK1) and LB medium (CK2) used as controls. After incubation for 48 hrs at 28 °C, 20 μL of 1 M NaOH solution was added before quantification live and dead J2s. Nematodes exhibiting body movement were categorized as alive, while those unresponsive were considered dead50. Each treatment was replicated six times, and the mortality of J2s was calculated as following: Relative mortality = (treatment mortality/control mortality)/(100-control mortality) × 100%.

Pot experiment of B. velezensis Y11.1
Two pot experiments, using normal and sterilized soil, were conducted in the greenhouse (China Agricultural University, Beijing, China, coordinates: 40°1′N, 116°16′E; 30/22 °C and a 14/10 hrs light/dark cycle at 60-70% humidity) from June to September 2022. Soil from a banana field in Nanning, Guangxi, China was characterized by a pH of 4.42 (in 1 mol/L KCl), and concentrations of organic matter, N, Olsen-P, and NH4OAc-K at 28.39 g/kg, 29.04 mg/kg, 3.78 mg/kg, and 117.27 mg/kg, respectively. Banana seedlings (Musa AAA Cavendish cv. Williams B6) were initially grown in vermiculite for 45 days before transplanting. For soil sterilization treatment, the soil was γ-irradiated (60Co-γ) with a maximum dose of 20 kGy for 24 h (Beijing Atomic High-tech Jinhui Radiation Technology Application Co. Ltd.). Seedlings were transplanted into 10 cm diameter plastic pots containing 0.5 kg of radiation-sterilized red soil. 50 mL of B. velezensis Y11.1 solution (106 CFU mL-1) were irrigated into the rhizosphere every 7 days, with 50 mL of water used as control. A total of 1000 Meloidogyne incognita second-stage juveniles (J2s, incubated at 28 °C within 3 days) were inoculated after the initial solution application, and plant and soil samples were collected 35 days post-inoculation. For normal soil treatment, seedlings were transplanted into 3 kg natural soil in 20 cm diameter plastic pots. Seven days post-transplantation, 300 mL of B. velezensis Y11.1 solution (106 CFU mL-1) were irrigated into the rhizosphere, with 3000 Meloidogyne incognita J2s inoculated simultaneously. B. velezensis Y11.1 solution was applied every 7 days, and 300 mL of water served as a control. Banana plants were harvested 60 days after J2s inoculation, and after measuring plant biomass, roots were carefully washed, and galls and egg masses from 10 g root sample were quantified. Soil nematodes were extracted and counted and then stored at -80 °C for root-knot nematode analysis. Each treatment consisted of four replicates.

Quantitative PCR of root-knot nematode
Soil nematode DNA from pot experiments under natural soil condition was extracted. The quality and concentration of DNA were assessed by 1.0% agarose gel electrophoresis and the NanoDrop® ND-2000 spectrophotometer (Thermo Scientific Inc., USA). The hypervariable region D2/D3 of the nematode 28S rDNA gene were amplified with root-knot nematode species-specific primer pairs RK28SF: 5’-CGGATAGAGTCGGCGTATC-3’ and MR: 5’-AACCGCTTCGGACTTCCACCAG-3’ with an ABI GeneAmp 7300 PCR thermocycler (ABI, CA, USA)51. The PCR reaction mixture consisted 4 μL 2x ChamQ SYBR Color qPCR Master Mix, 0.4 μL each primer (5 μM), 0.2 μL 50x ROX Reference Dye 1, 1 ng template DNA, and ddH2O to a final volume of 10 μL. The PCR amplification cycling conditions included an initial denaturation at 95 °C for 3 min, followed by 40 cycles of denaturing at 95 °C for 5 s, annealing at 58 °C for 30 s, and extension at 72 °C for 1 min, and with a final step at 4 °C. All samples were amplified in triplicate, and the specificity of the amplified fragments was confirmed by analyzing melt curves and electrophoresis on agarose gels. The results were expressed as copies/g soil by calculating the copy number of the target fragment from the standard curves.

Purification and quantitative assessment of bacillibactin production in B. velezensis Y11.1
Bacillibactin production by B. velezensis Y11.1 was assessed in iron-depleted liquid standard succinic medium (SSM) without additional iron (6 g L-1 K2HPO4, 3 g L-1 KH2PO4, 1 g L-1 (NH4)2SO4, 0.2 g L-1 MgSO4·7H2O, 4 g L-1 succinic acid, pH = 7.0) (Fig. 1c). After 48 hrs of incubation at 30 °C, cultures were diluted to OD600 = 0.8. Supernatant were collected following centrifugation (9400 g, 10 min at 4 °C) and filtration (using a 0.22 μm filter). Bacillibactin concentration was determined using a modified version of the chrome azurol S (CAS) assay52. Briefly, 4.3079 g of anhydrous piperazine was dissolved in 30 mL of water and adjusted the pH to 5.6 with HCl. Subsequently, 21.9 mg of HDTMA was dissolved in 50 mL water. Concurrently, a solution was prepared by mixing 1.5 mL of 1 mM FeCl3·6H2O (in 10 mM HCl) with 7.5 mL of 2 mM CAS. This solution was added to the HDTMA solution, and the mixture volume was adjusted to 100 mL with deionized water. One milliliter of supernatant was combined with an equal volume of CAS solution in a 2-mL centrifuge tube; deionized water served as a control. After 4 hrs of incubation at room temperature, after diluting the supernatants to w-fold, the OD630 of the supernatants (As) and deionized water (Ar) was measured by a plate reader (SpectraMax M5, Molecular Devices, Sunnyvale, USA). Bacillibactin-secreting capacity was quantified as Su = (Ar-As)/Ar. A calibration curve was established using desferrioxamine B (DFOB) to estimate bacillibactin concentration in B. velezensis Y11.1 supernatant (calibration curve equation: bacillibactin equivalent μM = (32.99 × Su + 0.2746) × 2, R2 = 0.999, p < 0.001).

The production and purification of bacillibactin were conducted following established protocol52. Briefly, B. velezensis Y11.1 was cultured in 10 L SSM medium at 30 °C and 180 r.p.m for 48 h. The culture media was centrifuged at 12000 g for 15 min and the resulting supernatant was filtrated. The culture medium was adjusted to pH 5.5 (using HCl) and adsorbed by XAD-4 resin. The bacillibactin-containing fraction was eluted using MeOH-H2O (1:1, v/v). The eluate was evaporated to dryness under vacuum. To enhance purity, the collected dry matter was subjected to a second adsorption by XAD-4 and elution with MeOD-H2O (1:1, v/v). The second eluate was concentrated to 5 mL, passed through a dextran gel (LH-20) column using MeOH-H2O 1:1 (v/v) as eluent to obtain the purified bacillibactin-containing fraction, which was then evaporated to dryness under vacuum.

Assessing the nematicidal activity of siderophore bacillibactin
To verify whether siderophore bacillibactin production affects nematode mortality, we cultured B. velezensis Y11.1 in iron-limited and iron-rich conditions. MKB medium (2.5 g l-1 K2HPO4, 2.5 g l-1 MgSO4·7H2O, 15 ml l-1 glycerin and 5.0 g l-1 casamino acids, pH 7.2) contains low quantities of residual iron but still allows B. velezensis Y11.1 to grow to a certain extent regardless of their ability to produce siderophores (iron-limited condition). As an iron-rich medium, the MKB medium was added with 50 μM FeCl3. Cultures were grown at 30 °C with shaking at 180 r.p.m 48 hrs. After centrifugation (4400 g, 5 min at 4 °C) and filtration (through 0.22 μm membrane), the cell-free supernatants were used to assayed the lethal effects of Meloidogyne J2s. MKB medium (used as control), iron-limited supernatant, and iron-rich supernatant were tested. a total of approximately 800 M. incognita J2s in a 1 mL suspension and 1 mL different supernatants were added into 24-well culture plates. After 48 hrs at 28 °C, 20 μL of 1 M NaOH solution was added before quantification live and dead J2s. The mortality rate was calculated as described above.

We next investigated the lethal test using purified bacillibactin on M. incognita and C. elegans. (1) The Meloidogyne juveniles (J2s) were incubated sterile water, dissolving the purified bacillibactin in sterile water to 100, 200, and 1000 μM. A volume of 500 μL Meloidogyne juveniles (containing approximately 100 J2s) and 500 μL various concentrations of bacillibactin were pipetted into 24-well cell culture plates. A volume of 1 mL sterile water was used as a control. The plates were then incubated at 25 °C for 48 hrs. Counting the dead nematodes as described above. (2) Age-synchronized C. elegans juveniles (18 hrs) were cultured in M9 medium (3 g/L KH2PO4, 6 g/L Na2HPO4, 5 g/L NaCl, 1 mL 1 M MgSO4) to a density of 100 juveniles per 500 μL. The purified bacillibactin were dissolved in M9 medium to 100, 200, and 1000 μM. A volume of 500 μL various concentrations of bacillibactin solutions and 500 μL C. elegans age-synchronized culture medium (containing approximately 100 J2s) were added into 24-well cell culture plates. M9 medium served as control. After incubating at 20 °C for 48 hrs, the dead nematodes were counted as described above.

Chemotaxis assay of bacillibactin with M. incognita and C. elegans
Chemotaxis assays were conducted to investigate the effects on the chemotactic motility of M. incognita and C. elegans under SSM medium and purified bacillibactin. Two lines were drawn 0.5 cm from the center at the bottom of 9 cm Petri dishes, and two points were drawn 1.0 cm from the lines and 1.5 cm from the center. A volume of 10 mL of 1% agar was added to petri dishes and allowed to dry. (1) For the chemotaxis assays of B. velezensis Y11.1 cultured in SSM medium on M. incognita, supernatant obtained as described above was used. The supernatant was diluted with SSM medium to concentrations of 0.01x, 0.1x, 0.5x, and 1x. Various concentrations of supernatant (50 μL) were placed on one point on the agar surface, while 50 μL of SSM medium was placed on the other point as a control. Approximately 100 M. incognita J2s (50 μL) were added to the center of the plate. After a 6-hour incubation at 28 °C, nematodes on the treated and control sides were counted. The chemotaxis index was calculated using the formula: chemotaxis index = (nematodes on the treated side - nematodes on the control side)/(nematodes on the treated side + nematodes on the control side). (2) For the chemotaxis assays of bacillibactin on M. incognita, the purified bacillibactin was diluted with sterile water to 50, 100, and 500 μM, 50 μL of various bacillibactin solutions were added to one point on the agar surface and 50 μL sterile water was added to other point, 50 μL M. incognita (approximately 100 J2s) was added on the center of the plate. Nematode were counted after a 6-hour incubation at 28 °C. (3) For the chemotaxis assays of bacillibactin on C. elegans, various concentrations of bacillibactin diluted with M9 medium (50, 100, and 500 μM) were tested. The next step of chemotaxis assays is described above. C. elegans were counted after a 6-hour incubation at 20 °C.

Pot experiment of bacillibactin
We identified the effect of bacillibactin in two pot experiments (normal and sterilized soil) from May to October 2024. The greenhouse, soil and banana seedlings were as described above. Seedlings were transplanted into 20 cm diameter plastic pots containing 3.0 kg of radiation-sterilized or natural red soil. A volume of 300 mL of bacillibactin solution (400 μM) were irrigated into the rhizosphere every 14 days, with 300 mL of water used as control. A total of 1800 M. incognita J2s were inoculated 7 days after the initial solution application. The plant and soil samples were collected 110 days post-inoculation. After measuring plant biomass, roots were carefully washed, and we counted the total galls and egg masses in the entire root system, and quantified the plant-parasitic nematodes abundance in 10 g root. Each treatment consisted of five replicates.

Statistical Analyses
Statistical analysis was conducted by using student t-test, two-way analysis of variance (ANOVA) with LSD as post-hoc test, as indicated in the Figure legends. A p-value < 0.05 was considered significant. The principal coordinate analysis (PCoA) and Bray-Curtis distance calculation were performed by R (v4.1.3) ‘phyloseq’ package (v.1.34.0). A co-occurrence network was constructed with all samples (two cropping times: Y2Y5, including samples from Y2 and Y5; Y8Y11, including samples from Y8 and Y11, each with 16 replicates). Our aim was to identify the ecological clusters of strongly associated ASVs, according to the methodology by Jiang et al.53. Indicator species analysis and likelihood ratio tests (LRTs) were employed pinpoint bacterial ASVs most significantly responsive to root-knot nematode suppression. Briefly, filtered communities were normalized using the BIOCONDUCTOR package EDGER, expressing normalized values as relative abundance counts per million (CPM)54. The R package INDICSPECIES was used to identify ASVs responsible for observed suppression effects through correlation-based indicator species analysis, with 104 permutations and significance set at p < 0.05. Differential ASV abundance between cropping times was tested using LRTs with R package EDGER, considering a false-discovery rate-adjusted value of p < 0.05 as indicative of responsiveness to cropping times. Indicator ASVs to cropping times were defined through both indicators species analysis and LRTs. Network modules, identified as substructures with a higher density of edges within groups than between them, were identified by calculating pairwise Spearman’s rank correlations between ASVs and using R package IGRAPH. Positive and significant correlations (Spearman’s ρ > 0.6, p < 0.01) and main modules in the network were visualized with the R package IGRAPH. Linear regression models were used to demonstrate the relationship between the relative abundance of the indicator modules and the relative abundance of Meloidogyne in the co-occurrence network. To identify the key microbial predictors of Meloidogyne abundance, Random Forest (RF) regression was performed using the R packages randomForest and rfPermute. Predictor variables consisted of the 40 most abundant ASVs from the two indicator modules (top 20 ASVs each from Module 1 and Module 2). The RF model was constructed using 500 regression trees (ntree = 500) with the importance = TRUE parameter enabled. The contribution of each ASV was quantified by the percentage increase in mean squared error (%IncMSE), and its statistical significance was determined through 1000 permutations (nrep = 1000). Plots including conceptual diagrams were assembled and aesthetically perfected by Adobe Illustrator CC, 2023 (Adobe, San Jose, USA).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Lu et al. investigated how long-term monoculture in banana fields leads to the development of suppressive soils against Meloidogyne spp., with a particular focus on the role of siderophore-producing Bacillus velezensis. The integration of field trials, microbial ecology analyses, and greenhouse experiments makes this a comprehensive and timely study. The findings offer valuable insights into microbiome-based strategies for the sustainable management of plant-parasitic nematodes (PPNs), aligning well with the goals of environmentally friendly agricultural development. This manuscript presents a well-executed and relevant study with strong potential impact. However, several key issues need to be addressed. I therefore recommend major revision. My detailed comments are as follows:\nMajor Comments\n1, The title suggests that soil suppressiveness is mediated by a cross-kingdom collaboration between free-living nematodes (FLNs) and siderophore-secreting Bacillus. However, the study presents only correlative evidence linking FLN abundance and Bacillus populations, without direct experimental demonstration of synergistic interaction. The title and related conclusions should be revised to more conservatively reflect the data and avoid overinterpretation.\n2, Sampling appears to have been conducted in years 1, 2, 4, 5, 7, 8, 10, and 11, omitting years 3, 6, and 9. The authors should clarify the rationale for this sampling scheme. Were there logistical, agronomic, or experimental reasons that precluded sampling in those years?\n3, Morphological assessments were performed for Y1, Y4, Y7, and Y10, whereas high-throughput sequencing was conducted for Y2, Y5, Y8, and Y11. Please clarify the reasoning behind this mismatched design and discuss any limitations it may introduce when interpreting inter-annual trends or correlations across datasets.\n4, While Figure 1 refers to morphological analyses, the full dataset is not included. Please either provide the complete morphological data in the main or supplementary materials, or indicate clearly where this information can be found.\n5, The term “sensitive ASVs” is introduced without definition or citation. Please clarify what is meant by “sensitive” in this context (e.g., responsiveness to environmental factors or abundance changes), and provide appropriate references if the term is established in the literature.\n6, The B. velezensis Y11.1 strain significantly reduced Meloidogyne populations in soil, but purified bacillibactin exhibited only modest nematicidal activity (11.5% mortality at 48 h). Given that the Y11.1 genome encodes other biosynthetic gene clusters, bacillibactin is unlikely to be the sole suppressive agent. Claims attributing suppressiveness exclusively to bacillibactin should be tempered, and the potential contribution of other metabolites should be discussed.\n7, The discussion introduces FLN diversity as a potential indicator of soil health (e.g., pp. 15 and 17), but this concept is not directly supported by the data presented and is peripheral to the study’s main objectives. These statements appear speculative and detract from the central focus. I recommend removing or significantly narrowing these sections to maintain clarity.\nMinor Comments\n1, Latin names such as Fusarium and Meloidogyne incognita should be italicized consistently throughout the manuscript. Similarly, gene names like 16S rRNA and 18S rRNA should follow standard formatting conventions. Please revise accordingly.\n2, Clearly and consistently refer to Meloidogyne incognita as the target PPN species throughout the manuscript to avoid ambiguity or confusion.", "reviewer_2": "This paper presents the results from experiments that endeavored to identify the cause of nematode suppression in a long-term banana mono-culture experiment.\nThere are several things that would improve this paper:\n1. People reading this study will certainly want to know if the reductions in nematodes in the long-term banana mono-culture experiment are associated with any beneficial effects to the plants. The authors should therefore provide data for both plant health and fruit yield for both the plants in mono-culture as well as the control plants.\n2. The title of the paper indicates that the authors have identified the cause of the putative nematode suppression. However, the data from the most important experiment, the Koch's postulates experiment (Figure 5f), does not support this claim:\na. In a suppressive soil, some microbe or some condition is inhibiting or killing the pathogen, and that microbe or condition is not transient. In the Koch's postulates experiment (Figure 5f), the authors applied a large number of their putatively suppressive bacterium (B. velezensis Y11.1) every 7 days for the 60 day experiment, which is 5 or 6 applications. If this bacterium was truly the cause of the putative nematode suppression in the long-term banana mono-culture experiment, the authors should not need to apply this bacterium every 7 days. The authors should therefore provide data from an experiment where the bacterium was applied once. As an aside, in a real agricultural situation, a biocontrol agent that requires application every 7 days would not be cost-effective.\nb. In addition, the authors added the nematodes and the putatively suppressive bacterium simultaneously in the Koch's postulates experiment. Since the authors showed that this bacterium killed the nematodes in in vitro experiments, the reduction in nematode numbers that the authors measured in this soil experiment could be an artifact of the simultaneous addition. It would be much better to add the bacterium and nematodes on different days to avoid this potential artifact.\nc. Also for the Koch's postulates experiment, since the authors are studying a root parasite, the authors should show the effect their bacterium has on the roots by providing dry root weight data. The authors should also provide dry shoot data.", "reviewer_3": "Key results: This manuscript presents a generally well-done study on the variation in soil microbiomes across distinct periods of continuous banana cropping, and their relationship to parasitic root knot nematode populations and banana root galls. As a research paper, there is value in the work. However, there are gaps in information and explanations of the data collection and/or analyses that should be addressed to strengthen the manuscript. Of biggest concern is the lack of any data provided to support the assertion that soil suppressiveness is `mediated by cross-kingdom collaboration between free-living nematodes and siderophore-secreting Bacillus’. The manuscript presents correlational data, but no specific mechanistic bases for supporting the thesis that this is a `cross-kingdom collaboration’. There is also not a clear definition of what `cross-kingdom collaboration’ means, other than that there are correlations between length of banana cultivation, population densities of free-living nematodes, siderophore-secreting Bacillus, and root knot infections. The abstract asserts that there are `synergistic roles’ of specific rhizobacterial genera and free-living nematodes, but does not provide either field or controlled environment data that supports synergistic interactions between rhizobacterial genera and free-living nematodes.\nValidity: Despite standing as a reasonable study of nematodes, soil microbes, and root knot infection, the broad assertion that the study makes about cross-kingdom collaboration is overambitious and not supported by the data presented in the paper.\nData, methodology, and analytical approaches:\n1. The methods need to provide much more detail and clarity on the field design. The fact that space is substituted for time in this study is a non-trivial detail that should be laid out clearly. Also, what were the dimensions of individual fields, the 10 plots within each field, and the total soil and root volumes collected? The authors state that each field had 10 replicates. By replicate, do you mean the 10 plots WITHIN each field? If so, those are not replicates, those are subsamples of the same field. It is critically important that this be addressed in the manuscript and the analyses. If true replicates, did you have 10 distinct fields (perhaps in a randomized complete block design?) for each cropping sequence? That does not seem to be what the work is describing, but if so, please make this very clear.\n2. The variable timing of the samples (March, July, October) also needs some clarity. Why 3 different sampling times? Were there 3 different sampling times in the same season, or did this differ between 2016 and 2017? Were the Y2 field samples collected in 2017 from the SAME field as that from which the Y1 samples were collected in 2016? Was this true across the 2016-2017 samples (the same field was sampled in 2 successive seasons), and did you perform any analyses comparing the 2016 and 2017 data for the same fields? These are significant gaps that need to be filled to strengthen the work and help the reader evaluate and contextualize the data.\n3. There is no information on the volume/biomass/area of roots assessed for root galls, or how this was done or what the replicates/experimental units were.\n4. The work includes both 16S and 18S sequencing. Is there a reason that you did not use the 18S data to provide information on the soil fungal biome? Also, the soil bacterial 16S data analyses are presented at a very broad taxonomic/phylogenetic level, despite the assertion in the methods that you made taxonomic assignments to individual ASVs. Were these ONLY at the phylum level? It seems like a significant gap not to dig more deeply into these data, as the distinct phyla reported in the work can have an enormous range of functional roles. More fine-scale taxonomic resolution of the 16S data would increase the value of these work.\n5. What is the relevance of the significant increases in bacterial density with longer-term banana cropping to root knot nematodes or gall incidence?\n6. Greater clarity in distinguishing between absolute and relative abundance is needed throughout the paper, especially when integrating distinct data types. For morphological assessments, presumably these numbers are all absolute abundances. However, for the sequencing data, these are only relative abundance metrics—how might this condition the strength of your interpretations? This needs to be called out in the text when reporting correlations (e.g. pp. 8 – 10).\n7. What is the evidence that free-living nematodes are actually beneficial in this system? You don’t present explicit data to justify this assertion for your samples or this system. There is no attempt to test this hypothesis in the controlled environment testing, unfortunately. This is a significant gap.\n8. Associations between the distinct nematode metrics (sequencing and morphological) should be addressed explicitly. Finally, you first use the abbreviation FLN on p. 4 in the manuscript, but do not define it. This should be formally defined at first use.\nClarity and context:\nA significant flaw in this work is the superficial presentation of the extensive published work on nematode suppressive soils for diverse cropping systems, including soils that have become nematode-suppressive following long-term crop monocultures. Similarly, there are not only many microbial taxa that are reported as being effective in nematode suppression, but also many Bacillus isolates that are documented to suppress nematodes. There are commercially-available Bacillus inoculants that are being sold as bionematicides. Collectively, the lack of information on this existing knowledge base on nematode-suppression is a substantial weakness of the manuscript. It seems that the justification of this gap in background information is that this work purports to take a mechanistic approach to understanding nematode suppression (bottom of p. 3, top of p. 4). However, I was not convinced that the data presented in this paper provide rigorous support for assertions of mechanistic understanding of cross-kingdom interactions in either the field or the controlled environment work. The discussion would benefit from including a bit of a skeptical eye to your results, specifically outlining potential weaknesses of the work and/or what needs to be done to further validate the assertions of mechanistic understanding. This would add some balance to the work.\nFinally, from a practical perspective, the manuscript would be improved by a brief consideration of other significant banana diseases, especially Fusarium wilt—which can interact significantly with nematodes/nematode colonization. I wondered if the fungivorous nematodes were having an impact on Fusarium wilt in the field and/or if this interacted in significant ways with the field data on gall numbers."}, "author_response": "Response 0: We sincerely thank the reviewers for their thoughtful and constructive\nfeedback on our manuscript (ID: NCOMMS-24-76527B). We have carefully revised the\nmanuscript in response to their comments and believe that the changes have\nsubstantially improved the clarity, rigor, and balance of the work. Specifically, we\naddressed three major areas of concern raised by the reviewers. First, clarifying that our\nfield data demonstrate correlations between free-living nematodes (FLNs) and Bacillus\nenrichment, while additional assays provided direct evidence of bacillibactin’s elective\neffects, which directly repels and kills root-knot nematodes, while attracting but not\nharming the FLN C. elegans, thereby distinguishing between correlation and causation;\nSecond, explaining the rationale and limitations of our sampling design, which\nemployed a space-for-time substitution approach, and providing additional\nmethodological details to ensure transparency; Third, reconciling the use of both\nmorphological and high-throughput sequencing approaches by demonstrating that,\ndespite their differences, they yielded consistent trends. In addition, we expanded the\nIntroduction and Discussion to better contextualize our findings within the extensive\nbody of research on nematode-suppressive soils and Bacillus-based biocontrol, and we\ntempered mechanistic claims to avoid overinterpretation. Finally, we have revised the\ntitle to “Soil suppressiveness to root-knot nematode in banana is associated with\nenrichment of free-living nematodes and siderophore-producing Bacillus under\ncontinuous cropping” which could accurately reflect the evidence presented.\nCollectively, these revisions not only resolve the reviewers’ concerns but also enhance\nthe novelty and broader significance of our study.\nSee below our point-by-point responses to all reviewers' comments.\nReviewer #1: (Remarks to the Author)\nLu et al. investigated how long-term monoculture in banana fields leads to the\ndevelopment of suppressive soils against Meloidogyne spp., with a particular focus on\nthe role of siderophore-producing Bacillus velezensis. The integration of field trials,\nmicrobial ecology analyses, and greenhouse experiments makes this a comprehensive and\ntimely study. The findings offer valuable insights into microbiome-based strategies for\nthe sustainable management of plant-parasitic nematodes (PPNs), aligning well with the\ngoals of environmentally friendly agricultural development. This manuscript presents a\nwell-executed and relevant study with strong potential impact. However, several key\nissues need to be addressed. I therefore recommend major revision. My detailed\ncomments are as follows:\nMajor Comments\nComment 1: The title suggests that soil suppressiveness is mediated by a cross-\nkingdom collaboration between free-living nematodes (FLNs) and siderophore-\nsecreting Bacillus. However, the study presents only correlative evidence linking FLN\nabundance and Bacillus populations, without direct experimental demonstration of\nsynergistic interaction. The title and related conclusions should be revised to more\nconservatively reflect the data and avoid overinterpretation.\nResponse 1: We thank the reviewer for this constructive comment. In accordance with\nthe suggestion, we refer to our Response 0 and have revised the title to “Soil\nsuppressiveness to root-knot nematode in banana is associated with enrichment of free-\nliving nematodes and siderophore-producing Bacillus under continuous cropping.” This\nmore accurately reflects the evidence presented. We also revised the Discussion (lines\n273-281) to highlight and address this valid point.\nWe agree that our field data primarily demonstrate correlations between FLN abundance\nand Bacillus enrichment. To strengthen this, we also performed validation experiments\nshowing that bacillibactin produced by B. velezensis Y11.1 directly repels and kills root-\nknot nematodes, while attracting but not harming the free-living nematode C. elegans\n(Fig. 6b, c). These results suggest a potential complementary role of siderophore-\nproducing Bacillus and FLNs in shaping suppressiveness, although we\nacknowledge that a direct mechanistic demonstration of synergistic interaction remains\nto be established.\nComment 2: Sampling appears to have been conducted in years 1, 2, 4, 5, 7, 8, 10, and\n11, omitting years 3, 6, and 9. The authors should clarify the rationale for this sampling\nscheme. Were there logistical, agronomic, or experimental reasons that precluded\nsampling in those years?\nResponse 2: We thank the reviewer for raising this point and refer to our Response 0 on\nthe major comments above. Specifically, our study used a space-for-time substitution\napproach to represent different durations of continuous banana cropping. In 2016, we\nsampled four groups of banana plantations that had been continuously cropped for 1 (Y1),\n4 (Y4), 7 (Y7), and 10 (Y10) years. In 2017, we re-sampled these same four groups\nplantations, which by then had reached 2 (Y2), 5 (Y5), 8 (Y8), and 11 (Y11) years of\ncontinuous cropping. Therefore, years 3, 6, and 9 were not omitted, but rather were not\nindependently sampled because our 2017 data represent the same sites one year older. This\ndesign was chosen because of the large scale of commercial banana plantations (hundreds\nof hectares each) and the logistical constraints of sampling multiple independent sites for\nevery cropping duration. By following the same fields over two years, we maintained\nspatial consistency and captured the temporal progression of both root-knot nematode\ndynamics and soil microbiome composition.\nWe have clarified this rationale in the revised Fig. 1a, Results text (lines 111–113) and\nMethods section (lines 417–424).\nComment 3: Morphological assessments were performed for Y1, Y4, Y7, and Y10,\nwhereas high-throughput sequencing was conducted for Y2, Y5, Y8, and Y11. Please\nclarify the reasoning behind this mismatched design and discuss any limitations it may\nintroduce when interpreting inter-annual trends or correlations across datasets.\nResponse 3: We thank the reviewer for this insightful comment. In the first year of\nsampling (2016; Y1, Y4, Y7, Y10), our primary goal was to identify overall patterns of\nroot-knot nematode infestation and soil nematode community composition; therefore, we\nused morphological identification as a cost-effective and well-established method.\nBased on these results, in the second year (2017; Y2, Y5, Y8, Y11), we applied high-\nthroughput sequencing to obtain higher-resolution profiles of nematode communities\nand, importantly, to simultaneously characterize microbial community composition\nfrom the same rhizosphere samples. This design allowed us to directly examine\npotential microbial–nematode associations and identify candidate microorganisms\ninvolved in disease suppression. Notably, both datasets converged on key findings—\nnamely, that Meloidogyne abundance declined in long-term continuous cropping soils,\nand that nematode community structure clearly separated short-term from long-term\ncropping stages (Fig. 3b,d and Fig. S1). We have revised Methods (lines 458-464) and\nDiscussion (lines 304-309) to clarify this rationale, and we now explicitly note the\nlimitations and complementarity of the two approaches.\nComment 4: While Figure 1 refers to morphological analyses, the full dataset is not\nincluded. Please either provide the complete morphological data in the main or\nsupplementary materials, or indicate clearly where this information can be found.\nResponse 4: The complete morphological data set is provided in Supplementary\nTables 2-4.\nComment 5: The term “sensitive ASVs” is introduced without definition or citation.\nPlease clarify what is meant by “sensitive” in this context (e.g., responsiveness to\nenvironmental factors or abundance changes), and provide appropriate references if the\nterm is established in the literature.\nResponse 5: We have defined “sensitive ASVs” in the Methods section (statistical\nanalyses), and cited Jiang et al. (Jiang et al., 2023). Briefly, sensitive ASVs refer to\nbacterial ASVs that were most significantly responsive to cropping duration. These were\nidentified using both indicators species analysis (R package INDICSPECIES) and\nlikelihood ratio tests (LRTs, R package EDGER) (Hartman et al., 2018; Jiang et al.,\n2023). To avoid confusion, we revised the term “sensitive ASVs” to “indicator ASVs”\nthroughout of the manuscript.\nHartman K, van der Heijden MGA, Wittwer RA, Banerjee S, Walser J-C,\nSchlaeppi K. 2018. Cropping practices manipulate abundance patterns of root\nand soil microbiome members paving the way to smart farming. Microbiome\n6(1).\nJiang Y, Wang Z, Liu Y, Han Y, Wang Y, Wang Q, Liu T. 2023. Nematodes and\ntheir bacterial prey improve phosphorus acquisition by wheat. New Phytologist\n237(3): 974-986.\nComment 6: The B. velezensis Y11.1 strain significantly reduced Meloidogyne\npopulations in soil, but purified bacillibactin exhibited only modest nematocidal activity\n(11.5% mortality at 48 h). Given that the Y11.1 genome encodes other biosynthetic gene\nclusters, bacillibactin is unlikely to be the sole suppressive agent. Claims attributing\nsuppressiveness exclusively to bacillibactin should be tempered, and the potential\ncontribution of other metabolites should be discussed.\nResponse 6: We agree that bacillibactin alone is unlikely to fully account for the\nsuppressive effect of B. velezensis Y11.1. Our intention is not to attribute\nsuppressiveness exclusively to bacillibactin but to highlight it as one demonstrable\nfactor with both repellent and lethal activity against Meloidogyne. We have therefore\nrevised the Discussion to temper our claims and to explicitly acknowledge the likelihood\nthat additional metabolites (e.g., lipopeptides, polyketides) may also contribute to\nsuppression (Vasantha-Srinivasan et al., 2024). We have also clarified that further work,\nsuch as mutant analyses of specific biosynthetic gene clusters, will be needed to\ndisentangle their relative roles. Accordingly, we have revised the Discussion section\n(lines 381-390).\nVasantha-Srinivasan P, Park KB, Kim KY, Jung WJ, Han YS. 2024. The role of\nBacillus species in the management of plant-parasitic nematodes. Front\nMicrobiol 15: 1510036.\nComment 7: The discussion introduces FLN diversity as a potential indicator of soil\nhealth (e.g., pp. 15 and 17), but this concept is not directly supported by the data\npresented and is peripheral to the study’s main objectives. These statements appear\nspeculative and detract from the central focus. I recommend removing or significantly\nnarrowing these sections to maintain clarity.\nResponse 7: We agree that positioning FLN diversity as a general indicator of soil health\nextended beyond the evidence provided. Accordingly, we have removed references to\n“soil health” and revised these passages to focus more narrowly on the observed\nassociations between specific FLN groups and Meloidogyne prevalence. We now\nemphasize that while some FLN groups were enriched in suppressive soils, their direct\nfunctional role in nematode suppression or plant growth was not tested here. We have\nadded a brief statement outlining future directions, such as controlled inoculation\nexperiments, to clarify that this remains a hypothesis for further investigation. Changes\nare included in lines 321-326 and line 407.\nMinor Comments\nComment 8: Latin names such as Fusarium and Meloidogyne incognita should be\nitalicized consistently throughout the manuscript. Similarly, gene names like 16S rRNA\nand 18S rRNA should follow standard formatting conventions. Please revise\naccordingly.\nResponse 8: We have thoroughly checked the entire manuscript and revised all Latin\nnames and standardized the formatting of gene names.\nComment 9: Clearly and consistently refer to Meloidogyne incognita as the target PPN\nspecies throughout the manuscript to avoid ambiguity or confusion.\nResponse 9: We focused on Meloidogyne spp. in both the field experiment and the pot\nexperiment using natural soil. For the in vivo and pot experiments conducted with\nsterilized soil, only M. incognita was used as the tested nematode. This clarification has\nnow been incorporated throughout the manuscript as requested.\nReviewer #2: (Remarks to the Author)\nThis paper presents the results from experiments that endeavored to identify the cause\nof nematode suppression in a long-term banana mono-culture experiment. There are\nseveral things that would improve this paper:\nComment 10: People reading this study will certainly want to know if the reductions in\nnematodes in the long-term banana mono-culture experiment are associated with any\nbeneficial effects to the plants. The authors should therefore provide data for both plant\nhealth and fruit yield for both the plants in mono-culture as well as the control plants.\nResponse 10: Thank you for this important comment. Our study was specifically designed\nto investigate the dynamics of nematode communities under continuous banana\nmonoculture, from the first year of planting (Y1) to the eleventh year (Y11). Because all\nplantations in this production region are monocultures, it was not possible to include a\n“non-monoculture control”. We agree that fruit yield is a useful indicator of plant\nperformance; however, banana yield under continuous monoculture is strongly constrained\nby the crop’s intrinsic characteristics and management practices. For this reason, yield data\nwere not collected. We have clarified this limitation in the revised manuscript (lines 314-\n320) and emphasized that the nematode community data provide an indirect but\ninformative proxy for plant health under long-term monoculture.\nComment 11: The title of the paper indicates that the authors have identified the cause of\nthe putative nematode suppression. However, the data from the most important\nexperiment, the Koch's postulates experiment (Figure 5f), does not support this claim:\nResponse 11: We thank the reviewer for raising this important point. Please see our\nResponse 0 to the major comments and the specific Response 1 on this point given to\nreviewer 1 above. To avoid potential misinterpretation, we have clarified the description\nof Fig. 5f in the revised manuscript (lines 219-223).\nComment 12a: In a suppressive soil, some microbe or some condition is inhibiting or\nkilling the pathogen, and that microbe or condition is not transient. In the Koch's postulates\nexperiment (Figure 5f), the authors applied a large number of their putatively suppressive\nbacterium (B. velezensis Y11.1) every 7 days for the 60 day experiment, which is 5 or 6\napplications. If this bacterium was truly the cause of the putative\nnematode suppression in the long-term banana mono-culture experiment, the authors\nshould not need to apply this bacterium every 7 days. The authors should therefore\nprovide data from an experiment where the bacterium was applied once. As an aside, in\na real agricultural situation, a biocontrol agent that requires application every 7 days\nwould not be cost-effective.\nResponse 12a: We agree that weekly applications of a biocontrol agent are not practical\nor cost-effective for real agricultural deployment. Our experiment was designed as a proof-\nof-concept assay to demonstrate the suppressive potential of B. velezensis Y11.1 under\ncontrolled conditions, rather than to optimize application strategies for the field. Because\nmicrobiome profiling does not provide absolute counts of Bacillus, and because the\nestablishment of effective populations may depend on interactions with co-occurring\nmicroorganisms, we applied Y11.1 repeatedly at relatively high densities to ensure its\nactivity was adequately tested. Similar repeated-application approaches are common in\nexperimental biocontrol studies (Doherty & Roberts, 2023; Hansel et al., 2024). We agree\nthat the next step will be to determine whether single or reduced-frequency applications\ncan provide comparable suppression in field settings. This has been included in our revised\nDiscussion section (lines 351-357).\nDoherty JR, Roberts JA. 2023. Topdressing biochar compost mixtures and biological\ncontrol organism applications suppress foliar pathogens in creeping bentgrass\nfairway turf. Plant Disease 107(8): 2346-2351.\nHansel J, Saville AC, Ristaino JB. 2024. Evaluation of a formulation of Bacillus\nsubtilis for control of Phytophthora blight of bell pepper. Plant Disease 108(4):\n1014-1024.\nComment 12b: In addition, the authors added the nematodes and the putatively\nsuppressive bacterium simultaneously in the Koch's postulates experiment. Since the\nauthors showed that this bacterium killed the nematodes in in vitro experiments, the\nreduction in nematode numbers that the authors measured in this soil experiment could\nbe an artifact of the simultaneous addition. It would be much better to add the bacterium\nand nematodes on different days to avoid this potential artifact.\nResponse 12b: We appreciate the reviewers concern regarding potential artifacts from\n’\nsimultaneous inoculation. Our intention was to approximate the natural condition in\nsuppressive soils, where root-knot nematodes and resident microbial antagonists interact\nin the rhizosphere concurrently. For this reason, we added B. velezensis Y11.1 and\nnematodes at the same time. This design allowed us to assess whether the bacterium could\nreduce nematode establishment and population growth under conditions that most closely\nreflect field conditions.\nComment 12c: Also for the Koch's postulates experiment, since the authors are studying\na root parasite, the authors should show the effect their bacterium has on the roots by\nproviding dry root weight data. The authors should also provide dry shoot data. Response\n12c: As requested, we have added dry root weight and dry shoot weight data for the Koch’s\npostulates experiment. These results are now included in Supplementary Fig. 6 (lines 212-\n213 and 216-217) and Supplementary Fig. 9 (line 253).\nReviewer #3: (Remarks to the Author)\nKey results: This manuscript presents a generally well-done study on the variation in soil\nmicrobiomes across distinct periods of continuous banana cropping, and their relationship\nto parasitic root knot nematode populations and banana root galls. As a research paper, there\nis value in the work. However, there are gaps in information and explanations of the\ndata collection and/or analyses that should be addressed to strengthen the manuscript. Of\nbiggest concern is the lack of any data provided to support the assertion that soil\nsuppressiveness is `mediated by cross-kingdom collaboration between free-living\nnematodes and siderophore-secreting Bacillus’. The manuscript presents correlational\ndata, but no specific mechanistic bases for supporting the thesis that this is a `cross-\nkingdom collaboration’. There is also not a clear definition of what `cross-kingdom\ncollaboration’ means, other than that there are correlations between length of banana\ncultivation, population densities of free-living nematodes, siderophore-secreting Bacillus,\nand root knot infections. The abstract asserts that there are ‘synergistic roles’ of specific\nrhizobacterial genera and free-living nematodes, but does not provide either field or\ncontrolled environment data that supports synergistic interactions between rhizobacterial\ngenera and free-living nematodes.\nValidity: Despite standing as a reasonable study of nematodes, soil microbes, and root\nknot infection, the broad assertion that the study makes about cross-kingdom\ncollaboration is overambitious and not supported by the data presented in the paper.\nResponse 13: We thank the reviewer for highlighting this important point. We\nacknowledge that our original use of the term cross-kingdom collaboration was not\n“ ”\nsufficiently addressed, overstating the strength of our evidence. This has now been\naddressed in the revised manuscript title, text (see lines 37-39, lines 284-285 and lines\n390-395) and Fig. 7. Please see our Response 0 to the major comments and the specific\nResponse 1 on this point given to reviewer 1 above for more details.\nData, methodology, and analytical approaches:\nComment 14: The methods need to provide much more detail and clarity on the field\ndesign. The fact that space is substituted for time in this study is a non-trivial detail that\nshould be laid out clearly. Also, what were the dimensions of individual fields, the 10\nplots within each field, and the total soil and root volumes collected? The authors state\nthat each field had 10 replicates. By replicate, do you mean the 10 plots WITHIN each\nfield? If so, those are not replicates, those are subsamples of the same field. It is critically\nimportant that this be addressed in the manuscript and the analyses. If true replicates,\ndid you have 10 distinct fields (perhaps in a randomized complete block design?) for\neach cropping sequence? That does not seem to be what the work is describing, but if\nso, please make this very clear.\nResponse 14: We have revised the Methods to clearly explain the spatial structure of\nsampling and the rationale behind the space-for-time substitution approach, please refer\nto our Response 2. In our study, we employed a space-for-time substitution approach\nacross banana plantations with different continuous cropping durations (from 1 to 11\nyears). For each cropping duration, we randomly selected 10 plantations for sampling. In\neach plantation, five banana plants of uniform growth status were selected using a five-\npoint (diagonal and center) sampling strategy, and their rhizosphere soils and roots were\npooled into a composite sample. Thus, each cropping duration yielded 10 biological\nsamples. These changes are now addressed in the Methods section (lines 425437).\nComment 15: The variable timing of the samples (March, July, October) also needs\nsome clarity. Why 3 different sampling times? Were there 3 different sampling times in\nthe same season or did this differ between 2016 and 2017? (Were the Y2 field samples\ncollected in 2017 from the SAME field as that from which the Y1 samples were\ncollected in 2016? Was this true across the 2016-2017 samples (the same field was\nsampled in 2 successive seasons), and did you perform any analyses comparing the 2016\nand 2017 data for the same fields? These are significant gaps that need to be filled to\nstrengthen the work and help the reader evaluate and contextualize the data.\nResponse: We thank the reviewer for this detailed and insightful comment. Below, we\nhave listed the individual criticisms and discussed them one by one.\nResponse 15a: Purpose of seasonal sampling:\nWe sampled across three time points (March, July, October) to capture the seasonal\ndynamics of root-knot nematode (RKN) disease and associated soil communities,\nintended to account for potential climatic variation in nematode behavior.\nResponse 15b: Sampling year and field consistency:\nYes, the Y2 field sampled in 2017 corresponded to the same field as the Y1 field sampled\nin 2016, and this was true for all pairs across the time series (e.g., Y1/Y2, Y4/Y5, Y7/Y8,\nY10/Y11). These pairs represent the same physical locations resampled one year later to\ntrack disease incidence and community shifts. We have revised and improved the Fig. 1a\nwhich could sample year and field consistency clearly.\nResponse 15c: Analysis approach and data presentation decisions:\nInitially, we analyzed disease incidence and community composition separately for each\nof the three sampling times per year (Fig. R1). However, we found no consistent or\nsignificant seasonal trends that altered the primary conclusions of the study. As a result,\nwe summarized disease incidence as mean values across the three sampling points. We\nhave revised the Methods section (see Response 14 for more details).\nFig. R1. The occurrence of root-knot nematode disease during continuous banana\ncultivation. a-b: disease incidence (a) and gall index (b) in banana orchards cultivated\ncontinuously for 1 year (Y1), 2 years (Y2), 4 years (Y4), 5 years (Y5), 7 years (Y7), 8\nyears (Y8), 10 years (Y10) and 11 years (Y11). Data presented as mean ± SE (n = 10).\nStatistical differences indicated by distinct lowercase letters (p ≤ 0.05) following\nDuncan’s test subsequent to one-way ANOVA. The small letters, capital letters, and\nsmall letters with punctuation reflected the statistical differences of spring, summer, and\nautumn, respectively.\nComment 16: There is no information on the volume/biomass/area of roots assessed\nfor root galls, or how this was done or what the replicates/experimental units were.\nResponse 16: Thank you for your valuable comment.\nIn the field experiment:\nApproximately 50 g of root samples were collected per banana plant. Galling of banana\nroot was rated using a 0–5 rating scale according to previous research, with some\nmodification (Bridge & Page, 1980). Thus, each cropping duration (e.g., Y1, Y2, etc.)\nwas represented by 10 biological replicates, each corresponding to a composite score\nfrom five plants in one plantation (Fig. 2c).\nIn the greenhouse pot experiments:\nFor the B. velezensis Y11.1 treatment, gall numbers were counted on ~10 g of fine root\ntissue per plant. For the bacillibactin treatment, we assessed the entire root system per\nplant to count total galls.\nThese methodological details have now been added to the revised Methods section\n(lines 438-442, 569-570 and 685–686) for better transparency.\nBridge J, Page SLJ. 1980. Estimation of root-knot nematode infestation levels on roots\nusing a rating chart. Tropical Pest Management 26(3): 296-298.\nComment 17: The work includes both 16S and 18S sequencing. Is there a reason that you\ndid not use the 18S data to provide information on the soil fungal biome? Also, the soil\nbacterial 16S data analyses are presented at a very broad taxonomic/phylogenetic level,\ndespite the assertion in the methods that you made taxonomic assignments to individual\nASVs. Were these ONLY at the phylum level? It seems like a significant gap not to dig\nmore deeply into these data, as the distinct phyla reported in the work can have an\nenormous range of functional roles. More fine-scale taxonomic resolution of the 16S data\nwould increase the value of these work.\nResponse 17: We thank the reviewer for this important comment regarding the\ntaxonomic depth of our microbial community analyses.\nResponse 17a: On the use of 18S data for fungal community analysis:\nWe chose 18S rRNA sequencing to capture soil nematode diversity, using nematode-\nspecific primers (NF1F/NR). These primers were selected and optimized for amplification\nof nematode taxa, not for comprehensive eukaryotic or fungal community profiling. We\nnow explicitly clarify this in the Methods section (lines 477-479). As we acknowledge the\nsignificant contributions of diverse soil organisms to the formation of nematode-\nsuppressive soils, we have offered a balanced discussion (lines 342-345) to acknowledge\nthis limitation and highlight it as a direction for future work.\nResponse 17b: On the resolution of the 16S data:\nIndeed, taxonomic classification was performed at the ASV level, and many ASVs were\nannotated to genus or species level. However, for initial data exploration and network\nmodularity analyses, we focused on phylum- (Fig. 4f), genus-level (Supplementary\nTable 1), and ASV-level (Supplementary Data 1) summaries to identify broad\ncompositional patterns and major responsive clades (lines 173-174). We have now\nexpanded Supplementary Tables to include full ASV (Supplementary Data 2) and\nmentioned the indicator ASVs in each module (Supplementary Data 1) in the main text,\nsuch ASVs-level taxonomy down to genus/species, where possible. We are grateful to\nthe reviewer for pointing out this opportunity to improve the taxonomic depth and\ntransparency of our data analysis.\nComment 18: What is the relevance of the significant increases in bacterial density with\nlonger-term banana cropping to root knot nematodes or gall incidence?\nResponse 18: In our dataset, total bacterial 16S rRNA gene copy numbers significantly\nincreased with longer durations of continuous banana cropping (Fig. 4b). However, the\ncorrelation analysis showed no statistically significant linear relationship between total\nbacterial density and gall index, disease incidence, or Meloidogyne relative abundance\n(Fig. R2). We believe this lack of correlation may be due to: (1) the fact that total bacterial\nabundance is a coarse metric that includes both beneficial and\nneutral/antagonistic/deleterious taxa; (2) the possibility that functional composition,\nrather than overall density, plays a more important role in suppressiveness. This valid\npoint is now addressed in the Discussion section (lines 332-336).\nFig. R2. Pearson correlation analysis between the total bacterial density and gall index\n(a), disease incidence (b), or Meloidogyne relative abundance (c). Shading associated\nwith lines in the panel indicates a 95% confidence interval.\nComment 19: Greater clarity in distinguishing between absolute and relative abundance\nis needed throughout the paper, especially when integrating distinct data types. For\nmorphological assessments, presumably these numbers are all absolute abundances.\nHowever, for the sequencing data, these are only relative abundance metrics—how\nmight this condition the strength of your interpretations? This needs to be called out in\nthe text when reporting correlations (e.g. pp. 8 – 10).\nResponse 19: We have carefully reviewed the manuscript and ensured that all instances of\n\"relative abundance\" and \"absolute abundance\" are clearly distinguished throughout the\ntext and figure legends. For both the morphological assessments (Fig. 3) and 18S\nrRNA sequencing (Fig. S1, Fig. 4g), we report relative abundance values — i.e., the\nproportion of a taxonomic group relative to total nematodes. Only in the case of total\nnematode counts (Fig. 2d) and quantitative PCR assays for Meloidogyne (Fig. 5f), we\nquantify absolute abundance (number of individuals or gene copies per gram of soil).\nWe have now added explicit caveats and clarifying language in the Results and\nDiscussion (lines 304-309) to acknowledge this limitation and encourage cautious\ninterpretation.\nComment 20: What is the evidence that free-living nematodes are actually beneficial\nin this system? You don’t present explicit data to justify this assertion for your samples\nor this system. There is no attempt to test this hypothesis in the controlled environment\ntesting, unfortunately. This is a significant gap.\nResponse 20: We agree that the results of the field experiments do not directly\ndemonstrate a functional or causal role of free-living nematodes (FLNs) in promoting\nplant health or suppressing root-knot nematodes in this banana system. Our conclusion\nthat FLNs are \"beneficial\" was based on their known ecological functions in other\nsystems and on the observed inverse correlation between FLN group abundance\n(particularly fungivores and omnivore-predators) and Meloidogyne relative abundance\nin our field samples (Fig. 3e). However, as the reviewer points out this correlation is not\nsufficient to assert benefit or causality, we revised the text throughout the manuscript to:\nMore accurately describe FLNs as \"putatively beneficial\" or \"functionally associated\nwith disease-suppressive soils\", and explicitly state that direct functional roles were not\ntested in this study. In support of their potential role, we cite extensive literature showing\nthat FLNs: Contribute to nutrient mineralization and microbial turnover, enhancing soil\nfertility (Ranoarisoa et al., 2020; Jiang et al., 2023). Directly suppress plant pathogens\nvia predation or antagonism, particularly through fungivorous (e.g., Aphelenchus,\nAphelenchoides) and omnivorous-predatory taxa (Wolfarth et al., 2013; Zhang et al.,\n2023). We now acknowledge this in the Discussion (lines 321-326).\nJiang Y, Wang Z, Liu Y, Han Y, Wang Y, Wang Q, Liu T. 2023. Nematodes and\ntheir bacterial prey improve phosphorus acquisition by wheat. New Phytologist\n237(3): 974-986.\nRanoarisoa MP, Trap J, Pablo AL, Dezette D, Plassard C. 2020. Micro-food web\ninteractions involving bacteria, nematodes, and mycorrhiza enhance tree P\nnutrition in a high P-sorbing soil amended with phytate. Soil Biology &\nBiochemistry 143.\nWolfarth F, Schrader S, Oldenburg E, Weinert J. 2013. Nematode–collembolan-\ninteraction promotes the degradation of Fusarium biomass and deoxynivalenol\naccording to soil texture. Soil Biology and Biochemistry 57: 903-910.\nZhang P, Zhang WJ, flu SJ. 2023. Fungivorous nematode and collembola alleviate\ndamping-off disease caused by in tomato. Plant and Soil 482(1-2): 175-189.\nComment 21: Associations between the distinct nematode metrics (sequencing and\nmorphological) should be addressed explicitly. Finally, you first use the abbreviation\nFLN on p. 4 in the manuscript, but do not define it. This should be formally defined at\nfirst use.\nResponse 21: Please see Response 3 for more details on the relationship between\nmorphological and sequencing-based nematode data. The term “FLN” (free-living\nnematodes) has now been formally defined upon first use in the revised manuscript\n(lines 75-76).\nClarity and context:\nA significant flaw in this work is the superficial presentation of the extensive published\nwork on nematode suppressive soils for diverse cropping systems, including soils that\nhave become nematode-suppressive following long-term crop monocultures. Similarly,\nthere are not only many microbial taxa that are reported as being effective in nematode\nsuppression, but also many Bacillus isolates that are documented to suppress nematodes.\nThere are commercially-available Bacillus inoculants that are being sold as\nbionematicides. Collectively, the lack of information on this existing knowledge base on\nnematode-suppression is a substantial weakness of the manuscript. It seems that the\njustification of this gap in background information is that this work purports to take a\nmechanistic approach to understanding nematode suppression (bottom of p. 3, top of p.\n4). However, I was not convinced that the data presented in this paper provide rigorous\nsupport for assertions of mechanistic understanding of cross-kingdom interactions in\neither the field or the controlled environment work. The discussion would benefit from\nincluding a bit of a skeptical eye to your results, specifically outlining potential\nweaknesses of the work and/or what needs to be done to further validate the assertions of\nmechanistic understanding. This would add some balance to the work.\nResponse 22: We sincerely thank the reviewer for this critical and constructive\nfeedback, which has helped us recognize several important areas for improvement in\nboth the literature framing and the interpretive balance of our manuscript. Below, we\nhave listed the individual criticisms and discussed them one by one.\nResponse 22a: Expanded background on nematode-suppressive soils and Bacillus-\nbased biocontrol:\nWe agree that our original Introduction and Discussion sections did not sufficiently\nreflect the rich body of research on nematode-suppressive soils. In the revised\nmanuscript, we have now changed the Introduction (lines 62-66) and the Discussion\n(lines 360-361):\nResponse 22b: Novelty of our work within this context\nWe fully acknowledge that Bacillus spp. are well established as nematode antagonists.\nOur study does not claim novelty in demonstrating that Bacillus can suppress root-knot\nnematodes. Instead, the novelty lies in: under long-term banana continuous cropping,\nBacillus species emerged as dominant taxa associated with soil suppressiveness, with\nBacillus velezensis Y11.1 identified as a representative/indicator strain enriched in\nsuppressive soils. Plate assays and pot experiments confirmed that suppression was\nmediated by secondary metabolites, in part by siderophores, which inhibited\nMeloidogyne. These findings suggest that while the nematicidal role of Bacillus is\nknown, continuous monoculture can selectively enrich specific Bacillus populations,\nwith secondary metabolite–mediated interactions providing a plausible mechanism for\ndisease suppression.\nResponse 22c: Reframing of mechanistic claims and acknowledgement of limitations\nWe agree with the reviewer that our original wording overstated mechanistic certainty.\nWe have now revised statements in the Abstract and Discussion to describe our findings\nas “evidence of association and potential functional contribution” rather than definitive\n“mechanistic proof”.\nFinally, from a practical perspective, the manuscript would be improved by a brief\nconsideration of other significant banana diseases, especially Fusarium wilt—which\ncan interact significantly with nematodes/nematode colonization. I wondered if the\nfungivorous nematodes were having an impact on Fusarium wilt in the field and/or if\nthis interacted in significant ways with the field data on gall numbers.\nResponse 23: We agree that Fusarium wilt, caused by Fusarium oxysporum f. sp.\ncubense (Foc), is a major threat to banana production worldwide and that interactions\nbetween Foc, nematodes, and other soil biota are important in shaping disease outcomes.\nPrevious studies indeed suggest that fungivorous nematodes can reduce fungal inoculum\nlevels in soil (Wolfarth et al., 2013; Zhang et al., 2023). However, the focus of our study\nwas the development of soil suppressiveness against root-knot nematodes. We therefore\ndid not systematically assess Fusarium wilt incidence in our field sites, as simultaneously\nmonitoring two major pathosystems would have introduced additional complexity beyond\nthe scope of this work. To acknowledge the reviewer’s point without diverting focus, we\nhave added a brief statement in the Conclusion highlighting that similar ecological\nprinciples may also be relevant to other banana pathogens such as Fusarium wilt, which\nwarrants future investigation. To address this, we have now mentioned in the Conclusion\n(lines 401-404).\nWolfarth F, Schrader S, Oldenburg E, Weinert J. 2013. Nematode–collembolan-\ninteraction promotes the degradation of Fusarium biomass and deoxynivalenol\naccording to soil texture. Soil Biology and Biochemistry 57: 903-910.\nZhang P, Zhang WJ, flu SJ. 2023. Fungivorous nematode and collembola alleviate\ndamping-off disease caused by in tomato. Plant and Soil 482(1-2): 175-189.\nPoint-by-point responses to the Reviewers' Comments:\nWe thank both reviewers for their valuable feedback, which has helped improve the\nclarity, rigor, and impact of the manuscript."}, {"round": 2, "reviewer_comments": {"reviewer_1": "I have carefully read the revised manuscript, and the authors have addressed my concerns.This paper should be a standout contribution to the field.", "reviewer_2": "The authors have provided adequate responses to the reviewer comments. However, the evidence supporting a causal link between Bacillus velezensis and nematode suppression remains weak."}, "author_response": "Point-by-point responses to the Reviewers' Comments:\nWe thank both reviewers for their valuable feedback, which has helped improve the\nclarity, rigor, and impact of the manuscript.\nReviewer #1:\nComment 1: I have carefully read the revised manuscript, and the authors have\naddressed my concerns. This paper should be a standout contribution to the field.\nResponse 1: We are grateful to Reviewer #1 for the positive assessment and\nencouraging remarks. We appreciate the recognition that the revised manuscript\naddresses the reviewer’s concerns and contributes meaningfully to the field.\nReviewer #2:\nComment 2: The authors have provided adequate responses to the reviewer comments.\nHowever, the evidence supporting a causal link between Bacillus velezensis and\nnematode suppression remains weak.\nResponse 2: We thank Reviewer #2 for raising the important point regarding the causal\nlink between Bacillus velezensis and nematode suppression. We would like to\nemphasize that our conclusions are supported not only by consistent field associations,\nbut also by controlled experimental evidence. Specifically, indoor bioassays\ndemonstrated significant inhibitory and repellent effects of B. velezensis against root-\nknot nematodes. Further analysis identified the siderophore bacillibactin as an active\ncontributor to these effects, and greenhouse pot experiments confirmed that B.\nvelezensis application effectively reduced nematode infection and disease severity.\nWhile we acknowledge that additional work is needed to fully resolve mechanistic\ninteractions under field conditions, these complementary experimental results provide\nstrong functional evidence for a direct suppressive role of B. velezensis.\nWe sincerely thank the editors and both reviewers for their careful evaluation of our\nrevised manuscript and for the constructive and insightful comments."}]

Decision letter