Organic pollutant-induced long-distance ROS signaling drives plant systemic acquired acclimation via rhizomicrobiota

Li, Yong; Zhang, Kaiwei; Zhang, Hui; Li, Mei; Ma, Liya; Liu, Yang; Feng, Fayun; Jiang, Wayne; Ge, Jing; Chen, Jian; Yu, Xiangyang

doi:10.1038/s41467-025-64138-y

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Published: 13 October 2025

Organic pollutant-induced long-distance ROS signaling drives plant systemic acquired acclimation via rhizomicrobiota

Yong Li^1,2,
Kaiwei Zhang^1,2,
Hui Zhang^1,2,
Mei Li^1,2,
Liya Ma^1,2,
Yang Liu^1,2,
Fayun Feng^1,2,
Wayne Jiang³,
Jing Ge^1,2,
Jian Chen ORCID: orcid.org/0000-0002-0766-653X^1,2 &
…
Xiangyang Yu ORCID: orcid.org/0000-0002-2831-172X^1,2,4

Nature Communications volume 16, Article number: 9077 (2025) Cite this article

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Abstract

Plants can recruit beneficial rhizomicrobes to combat environmental stimuli, but the upstream signaling through which plants sense stress to initiate rhizomicrobial recruitment still remains unclear. This study elucidates the role of long-distance ROS signaling in driving the recruitment of beneficial rhizobacteria to establish systemic acclimation following local organic pollutant stress. Plant leaves sense various organic pollutants to generate ROS, followed by the occurrence of a long-distance ROS wave from leaves to roots via a Ca²⁺-RBOH-ROS signaling module. Elevated ROS in roots plays dual functions. First, ROS stimulates plant carbon release into the rhizosphere by increasing the permeability of root cell membranes. The released carbon flux enriches plant-beneficial bacterial genera, which in turn promotes plant growth and pollutant degradation. Second, NO acts downstream of ROS to loosen root cell walls, facilitating rhizobacterial colonization. Our findings show how plants deploy systemic signaling acquire help from rhizomicrobes, extending our understanding of plant environmental adaptability.

Introduction

Environmental pollution poses a severe threat to global ecosystem, becoming one of the most pressing challenges worldwide^1,2. Plants frequently encounter a variety of pollutants, including inorganic (e.g., salts and heavy metals) and organic pollutants (e.g., pesticides, polycyclic aromatic hydrocarbons, and polychlorinated biphenyls), which can significantly impair plant growth and development^3,4. In response, plants have evolved diverse strategies to adapt to pollutant exposure⁵. Besides intrinsic physiological adjustment, plants are good at seeking help from rhizosphere microbiota⁶. These recruited microbes can colonize inside of plants to bolster plant resistance and to degrade pollutants in vivo^7,8. Plant root exudates play an important role in reshaping rhizosphere microbial community and facilitating the recruitment of beneficial microbiota⁹.

Most of the studies focus on soil pollutants because of the direct interaction among plant roots, soil pollutants, and rhizosphere microbiomes. Recently, terrestrial plant health responses to air pollutants have been drawing great attention¹⁰. Plant leaves are frequently exposed to organic pollutants through pesticide spraying and atmospheric deposition of pollutants, often resulting in leaf injury^11,12. These organic pollutants can not only regulate plant metabolism but also affect plant microbiota community¹³. Studies have shown that the pesticide N-(3,5-dichlorophenyl) succinimide increases the abundance of Acinetobacter while decreasing the abundance of Stenotrophomonas and Flavobacterium in the tobacco phyllosphere¹⁴. Fungicide enostroburin has been observed to elevate the abundance of Pantoea in the wheat phyllosphere¹⁵. Our research has demonstrated that foliar exposure of pesticide imidacloprid could facilitate the recruitment of rhizosphere beneficial bacteria, which could help to compensate for pesticide-induced stress in plants^16,17. Small molecular organic acids in rice root exudates could enhance the colonization of endophytic Enterobacter cloacae TMX‑6, a thiamethoxam degrading bacterium¹⁸. These findings raise questions about how the rhizomicrobiome is altered by foliar sensing of stresses. The interactions between root exudates and rhizosphere microbiota have been well established¹⁹, but little is known about the involvement of upstream plant intrinsic signaling driving this process, especially regarding long-distance signaling transduction derived from locally stressed plant organs.

In plants, reactive oxygen species (ROS) are multifunctional long-distance signals, modulating plant adaptation to environmental stimuli, including pollutant stress. ROS can sense stress conditions and activate plant stress-response networks²⁰. Respiratory burst oxidase homolog (RBOH) proteins with NADPH oxidase activity are critical enzymes to produce ROS in plants under stress conditions. RBOH is considered as a regulatory hub connecting the interaction between ROS and other signaling. Ca²⁺, an ubiquitous long-distance messenger in plant cells, binds to the EF-hand motifs in ROBHs to activate them for ROS burst. And ROS can promote the flux of Ca²⁺ in neighboring cells to form a positive feedback loop, further achieving the cell-to-cell and organ-to-organ signaling transduction of ROS in plants^20,21. Local injury resulting from pathogen infection or pest attack is able to trigger systemic transmission of Ca²⁺ and ROS signaling to non-injured plant organs^22,23, driving us consider the possibility of similar signaling responses in plants upon local exposure of pollutants. In addition, nitric oxide (NO), an intrinsic gasotransmitter, acts as a vital player of ROS in plant stress responses²⁴. ROS can affect cellular NO concentrations by fine-tuning S-nitrosoglutathione reductase activity, thereby activating protection against oxidative stress²⁵. NO can regulate cellular ROS levels by enhancing the activity of antioxidant enzymes such as catalase and superoxide dismutase under stress conditions²⁶. Although direct evidence for the regulation of rhizosphere microbiota by these plant intrinsic signals is currently lacking, their roles in Legume-rhizobial symbiosis have been documented²⁷. These findings offer valuable insights and highlight the need for further exploration into the potential role of these signals in rhizomicrobiota recruitment.

A hypothesis is proposed that foliar exposure of pollutants might trigger the long-distance transmission of ROS signaling from leaves to roots, subsequently inducing the release of root exudates to recruit rhizosphere beneficial microbes for the establishment of plant resistance. In this study, this hypothesis was tested using Brassica rapa, a widely cultivated vegetable, which was exposed to a range of organic pollutants, including insecticide thiamethoxam, fungicide tebuconazole, herbicide acetochlor, polycyclic aromatic hydrocarbon phenanthrene, and polychlorinated biphenyl trichlorobiphenyl. This study aims to uncover a universal long-distance signaling mechanism that modulates the interaction between root exudates and rhizomicrobiota recruitment in response to localized foliar pollutant exposure.

Results

Foliar exposure to various organic pollutants enriches common beneficial bacteria in rhizosphere

The leaves of Brassica rapa were treated with five different organic pollutants, thiamethoxam (Thia), tebuconazole (Teb), acetochlor (Ace), phenanthrene (Phen), and trichlorobiphenyl (Trich) and water with 0.1% polysorbate-80 (as the control group). Plant roots and soil were shielded to prevent exposure to organic pollutants. The rhizosphere bacterial community for each treatment was analyzed two weeks post-exposure. The 16S rRNA amplicon sequencing yielded a total of 3,469,952 bacterial sequences with 8,164 bacterial OTU sequences. Principal coordinates analysis (PCoA) revealed that foliar exposure of organic pollutants was able to reshape the composition of rhizosphere bacterial communities (PERMANOVA, P < 0.05; Fig. 1a). Foliar exposure of organic pollutants resulted in significantly higher rhizosphere bacterial diversity than the control group, as indicated by Shannon index (two-sided Wilcoxon rank-sum test, P < 0.05, Fig. 1b). In total, 650 genera were identified across all samples, among which 138 exhibited significant changes in pollutant-treated groups compared to the control group (LEfSe with two-sided Wilcoxon rank-sum test, P < 0.05, LDA effect size > |2.5|; Supplementary Data 1). Among these genera, 37 to 48 exhibited significantly increases, while 27 to 39 showed decreases across the different pollutant groups compared to the control (Fig. 1c, d). Treatment with organic pollutants led to an increase in 19 genera, with Sphingomonas and Lysobacter exhibiting the highest average relative abundances among all samples (Fig. 1e, f).

**Fig. 1: Effects of leaf-exposure organic pollutants on plant rhizosphere bacterial communities and feedback of beneficial bacteria to plants.**

No residues of these organic pollutants or their key intermediates were detected in roots or soil (Supplementary Fig. 1), which rules out the possibility of direct stimulation of the rhizosphere environment by these organic pollutants or their intermediates. This indicates that different pollutants, acting as stress factors, indirectly induce changes in the rhizosphere bacterial community, and the reshaped microbial community may subsequently regulate plant health. Additionally, compared to Phen and Trich, exposure of plants to pesticides is more common in agricultural production due to its widespread use. We selected the enriched genera Sphingomonas and Lysobacter, along with Thia, as representatives to assess their effects on organic pollutant degradation and plant growth. Representative strains, Sphingomonas sp. LSS1 and Lysobacter sp. LSS2 were isolated from Thia-treated rhizosphere soil (Supplementary Fig. 2). The root of plant seedlings was separately inoculated with 1.0 mL of LSS1 (OD_600nm = 0.5), LSS2 (OD_600nm = 0.5), and their proportional synthetic microbial community (SynCom), while the control group was treated with 1.0 mL sterile water²⁸. Inoculation of roots with these strains for 20 days significantly increased plant biomass as compared to the control, with the greatest effect observed at the SynCom group (Fig. 1g, h). Following inoculation, plants were foliar-sprayed with Thia. Ten days post-treatment, SynCom-inoculated plants exhibited 25.2% to 55.1% lower concentrations of Thia compared to the control and single-strain groups (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 1i). In addition, SynCom showed the greatest degradation effect on Thia, with reductions in pollutant concentration of about 38.8% compared to controls, followed by Strain LSS1 (25.4%) (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 1j). No degradation effect was observed with Strain LSS2 (one-way ANOVA, two-sided Tukey’s test, P > 0.05). These results demonstrate that foliar exposure of different organic pollutants co-enriches rhizosphere beneficial bacteria, promoting pollutant degradation in vivo and plant growth.

Foliar exposure of organic pollutants promotes plant carbon effluxes

Root exudates serve as a crucial link in the interaction between plants and rhizosphere microbes⁹. In soil-plant systems, the composition of rhizosphere metabolites is relatively complex, encompassing root exudates, microbial metabolites, and soil components. To eliminate the influence of microbial metabolites and soil components, hydroponic conditions were employed to analyze the effect of organic pollutants on plant root exudates. The root exudate solutions of hydroponic plants, leaf-sprayed with different pollutants and water containing 0.1% polysorbate-80 (as the control group), were collected during 48 hours of exposure. The biomass of the dried root exudates significantly increased in organic pollutant groups, exhibiting 1.14–1.25 times higher values compared to the control group (two-sided Student’s t-test, P < 0.05; Fig. 2a). Metabolomic analysis using LC-QTOF/MS revealed that the metabolic profiles of root exudates were significant changed upon foliar exposure of organic pollutants (PCA, PERMANOVA, P < 0.05; Supplementary Fig. 3). A total of 262 metabolites showing significant changes were identified (two-sided Student’s t-test with P < 0.05, PLS-DA with VIP > 1; Supplementary Fig. 4, Supplementary Data 2). Among these metabolites, 239–259 metabolites were significantly increased in treatments with different organic pollutants, compared to the control group. A total of 215 metabolites increased consistently, accounting for 82.1% of total metabolites, despite some variation in the degree of increase (Fig. 2b, c). These metabolites mainly included primary and secondary metabolites, such as amino acids, organic acids, fatty acids, lysophosphatidylcholines, nucleosides, vitamins, phenolic acids, flavonoids, and alkaloids (Fig. 2d). These findings suggest that foliar exposure of different organic pollutants could consistently enhance the release of a large quantity of small-molecule organic carbon from roots.

**Fig. 2: Effect of various organic pollutants on plant root exudate secretion and preferential utilizability of rhizosphere-enriched bacterial strains on root exudate metabolites.**

Root carbon effluxes recruit rhizosphere beneficial bacteria

Correlation analysis revealed complex relationships between altered root exudates and co-enriched rhizosphere bacterial genera upon foliar exposure of organic pollutants (Spearman’s rank correlation, P < 0.01; Supplementary Fig. 5). A capillary assay demonstrated that both Sphingomonas sp. LSS1 and Lysobacter sp. LSS2 displayed strong chemotaxis toward root exudates from Thia-treated plants. The colony-forming units (CFUs) of Strain LSS1 and LSS2 in capillary tubes containing Thia-treated plant root exudates were 1.8–2.4 times higher than those with control root exudates (two-sided Student’s t-test, P < 0.05; Fig. 2e, f). Subsequently, root exudate solutions from plants exposed to Thia were collected to culture the bacterial strains in order to investigate the preferences of these two bacterial strains for root exudate metabolites. Exometabolomics analysis revealed that a total of 89 and 79 metabolites were down-regulated in the solution culturing Strain LSS1 and LSS2, respectively, compared to the solution without strains (two-sided Student’s t-test with P < 0.05, PLS-DA with VIP > 1; Fig. 2g; Supplementary Fig. 6, Supplementary Data 3). Among theses metabolites, 65 metabolites were commonly decreased, indicating likely bacterial consumption. Both strains showed a strong preference for amino acids, fatty acids, and nucleotides (Fig. 2g; Supplementary Fig. 7). Additionally, Strain LSS1 tended to consume small-molecule organic acids (citric acid, quinic acid, and pipecolic acid), while Strain LSS2 favored lysophosphatidylcholines and vitamins (riboflavin and pantothenic acid). These findings indicate that root carbon effluxes resulting from foliar exposure to organic pollutants play a role in the recruitment of rhizosphere beneficial bacteria.

Foliar exposure of organic pollutants triggers long-distance transmission of Ca²⁺-dependent ROS signaling from leaves to roots

Nitro-blue tetrazolium staining revealed that foliar exposure of Thia resulted in an increase in ROS contents in roots of B. rapa (Fig. 3a). To investigate the origin of root ROS, the spatial and temporal dynamics of ROS were analyzed throughout the plant following exposure to various organic pollutants. Foliar exposure of five tested organic pollutants could enhance ROS levels in both leaves and roots, exhibiting an increase of 1.18-1.47 and 1.34-1.81 times compared to those from the control group, respectively (two-sided Student’s t-test, P < 0.05; Fig. 3b). Thia was selected to study the time-dependent changes of ROS. ROS in leaves began to increase significantly at 3 h post-exposure, while ROS in stems and roots exhibited delayed responses, with significant increases beginning at 6 hours after foliar Thia treatment (two-sided Student’s t-test, P < 0.05; Fig. 3c). The endogenous Ca²⁺ level was measured because of the pivotal role of Ca²⁺ in facilitating ROS signaling transduction. While Ca²⁺ levels in leaves remained unchanged following Thia exposure, Ca²⁺ levels in stems and roots began to increase at 6 h post-treatment, aligning with the ROS pattern (two-sided Student’s t-test, P < 0.05; Fig. 3d). Inhibition of Ca²⁺ channels with La³⁺ in leaves blocked ROS accumulation in stems and roots after foliar exposure of Thia (two-sided Student’s t-test, P > 0.05; Fig. 3e). These results suggest that foliar exposure of organic pollutants triggers Ca²⁺-dependent ROS signaling transmission from leaves to stems and roots.

**Fig. 3: Intercellular transmission of Ca²⁺-induced ROS signaling from stressed plant leaves to roots.**

In B. rapa, the RBOH is responsible for ROS production in Ca²⁺-dependent ROS propagation. The RBOH gene family, comprising total of 14 members, has been identified in B. rapa²⁹. Foliar exposure of Thia for 6 h significantly upregulated the expression of 6, 4 and 4 BrRBOH members in leaves, stems, and roots, respectively, compared to the corresponding tissues from the pre-exposure group (two-sided Student’s t-test, P < 0.05; Figs. 3f, S8) and other detected BrRBOHs showed a slight decrease or no change in expression in different comparisons (Fig. 3f, Supplementary Fig. 8). Notably, among these upregulated genes, BrRBOH03 exhibited an increase of 4.18–6.28 times in the whole plants exposed to Thia, compared to the pre-exposure group (two-sided Student’s t-test, P < 0.05; Fig. 3f, Supplementary Fig. 8). The expression of BrRBOH07, BrRBOH08 and BrRBOH11 was additionally increased in the stems of Thia-treated plants and the expression of BrRBOH06, BrRBOH12, and BrRBOH13 was additionally increased in the roots of Thia-treated plants, exhibiting an increase of 1.75–7.78 times, compared to the corresponding tissues from the pre-exposure group (two-sided Student’s t-test, P < 0.05; Figs. 3f, S8). The activation of BrRBOH genes, accompanied by the accumulation of ROS and Ca²⁺ in roots, indicates the involvement of BrRBOHs in the long-distance transmission of ROS upon foliar treatment with organic pollutants.

ROS increase in roots enhances plant carbon efflux into rhizosphere

Increased intracellular ROS can lead to the increase in cell membrane permeability, as indicated by increased relative conductivity. Foliar exposure of the five tested organic pollutants resulted in significant increase at 1.32–1.68 times in root membrane permeability compared to the control group (two-sided Student’s t-test, P < 0.05; Fig. 4a). A time-course experiment suggested that root relative conductivity began to increase after foliar exposure of Thia at 6 h, paralleling the increase in root ROS content (Fig. 4b). The cell membrane regulates the exchange of materials between the inside and outside of cells. Therefore, we hypothesize that the rise in ROS leads to an increase in root membrane permeability, causing a higher efflux of plant carbon. To further verify this hypothesis, we sought to increase membrane permeability without enhancing ROS levels. Linoleic acid is one of the key components of the cell membrane lipids, which is able to increase membrane fluidity and permeability³⁰. Linoleic acid levels were significantly increased at 2.4–2.8 folds in root tissues from pollutant-treated groups compared to the control group (two-sided Student’s t-test, P < 0.05; Supplementary Fig. 9). As expected, roots treated with linoleic acid showed increased relative conductivity without altering endogenous ROS levels (Fig. 4c, d), suggesting that the addition of linoleic acid was able to mimic the enhancement of membrane permeability. Additionally, roots treated with ROS donor H₂O₂ similarly elevated endogenous ROS levels, resembling the effects observed in the Thia-treated group (Fig. 4e). The increase of ROS levels under H₂O₂- and Thia-treated groups was inhibited by further addition of N-acetyl-L-cysteine (NAC, ROS scavenger) (Fig. 4e).

**Fig. 4: Plant root exudate secretion regulated by ROS-induced cell membrane permeability.**

The biomass of the dried root exudates significantly increased in H₂O₂- and linoleic acid-treated groups, exhibiting 1.26 and 1.21 times higher values than that of the control group, respectively (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 4f). The increase in root exudates observed under H₂O₂- and linoleic acid treatment was comparable to that in the Thia-treated group (Fig. 2a). The increase of biomass of the dried root exudates under H₂O₂- and Thia-treated groups was also inhibited by adding NAC (Fig. 5f). Metabolomic analysis revealed that among detected 262 metabolites in the metabolites of B. rapa root exudates, 226 and 221 ones were significantly increased in H₂O₂- and linoleic acid-treated group, respectively, as compared to the control (two-sided Student’s t-test with P < 0.05, PLS-DA with VIP > 1; Fig. 4g, Supplementary Data 4). A cross-analysis identified 206 common metabolites that were induced by Thia, H₂O₂, and linoleic acid, accounting for ~80.8% of total increased metabolites (Fig. 4h). These findings demonstrate that elevated ROS in roots was able to enhance the release of plant carbons into the rhizosphere by increasing the permeability of root cell membranes.

**Fig. 5: Colonization of rhizosphere beneficial bacteria regulated by ROS-mediated NO.**

Root NO acts downstream of ROS to promote beneficial bacteria colonization

The above results demonstrated that long-distance ROS signaling was capable to regulate root exudates to recruit rhizosphere bacteria. Next, this study investigated how these bacteria colonized plant roots by examining the role of endogenous NO, a potential signaling molecule. Foliar exposure to each of five tested organic pollutants for 24 h significantly enhanced NO levels in roots, exhibiting an increase of 2.07–2.60 times compared to the control group (two-sided Student’s t-test, P < 0.05; Fig. 5a). Spatial and temporal analysis indicated that foliar exposure of organic pollutant (Thia) did not change endogenous NO level in leaves and stems (two-sided Student’s t-test, P > 0.05; Fig. 5b), suggesting that NO generation occurred specifically in roots rather than from shoots. This root-specific NO production was supported by increased nitrate reductase and nitric oxide synthase activity in roots following Thia exposure (two-sided Student’s t-test, P < 0.05; Fig. 5c, d). Furthermore, NO accumulation in roots lagged behind ROS accumulation, indicating NO might act as downstream of ROS in response to foliar pollutant exposure.

Pharmacological experiments were performed to clarify the relationship between ROS and NO in roots upon foliar Thia exposure. Foliar exposure to Thia or direct root exposure to H₂O₂ and sodium nitroprusside (SNP, NO donor) resulted in an increase in root NO content, but this increase was inhibited by the further addition of the ROS scavenger NAC and 2-(4-carboxyphenyl)−4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (c-PTIO, NO scavenger) (Fig. 5e). However, the addition of c-PTIO had little effect on root ROS levels when applied alongside Thia or SNP (Fig. 5f). These results suggest that elevated root ROS drives NO production in response to foliar pollutant exposure. Additionally, Thia and SNP applications enhanced the cellulase activity in roots, exhibiting an increase of 2.07 times compared to controls (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 5g). Reducing NO levels with cPTIO in both the Thia and SNP groups significantly decreased cellulase activity, making it comparable to that of control plants (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 5g). As cellulase enzymes hydrolyze cellulose within the cell walls, they facilitate cell wall relaxation. Calcofluor white staining revealed that cell wall integrity was compromised in both Thia- and SNP-treated roots, as observed under a fluorescence microscope (one-way ANOVA, two-sided Tukey’s test, P < 0.05; Fig. 5h, i). Reducing NO levels with cPTIO in both the Thia and SNP groups did not affect cell wall integrity, which was comparable to that of control plants (one-way ANOVA, two-sided Tukey’s test, P > 0.05; Fig. 5i).

We placed the plants treated with ROS and SNP, along with their inhibitor-treated counterparts, into a solution containing RFP-labeled LSS1 and LSS2 to investigate the effects of ROS and SNP on bacterial colonization behavior. Observation under a fluorescence inverted microscope revealed that the roots of plants treated with Thia, H₂O₂, and SNP showed more red fluorescence compared to the control, with the CFUs of the strains being ~1.85–2.59 times higher (Fig. 5j, k; Supplementary Fig. 10). The increase of colonization under Thia-, H₂O₂- and SNP-treated groups was inhibited by further adding NAC or cPTIO (Fig. 5k). Furthermore, neither Strain LSS1 nor LSS2 displayed chemotaxis towards H₂O₂, thereby eliminating the possibility of ROS playing a direct role in the recruitment and colonization of these bacterial species (Supplementary Fig. 11). Therefore, it can be concluded that ROS-mediated NO increase promotes cell wall relaxation via increasing cellulase activity and facilitates the colonization of rhizosphere beneficial bacteria.

Discussion

As sessile organisms, plants have evolved sophisticated strategies to protect themselves from environmental stressors, including reshaping the rhizosphere microbiome^31,32. However, very little is known about how plants sense pollutant stress to initiate the recruitment of beneficial rhizosphere microbes. In the present study, four lines of evidence suggest that long-distance transmission of ROS signaling is key in sensing foliar exposure of organic pollutants to facilitate the recruitment and colonization of beneficial rhizobacteria in roots. Initially, foliar exposure of organic pollutants resulted in local ROS burst, followed by long-distance signaling transmission to roots through Ca²⁺-ROBH-ROS signaling cascade. Secondly, ROS activation in roots increased membrane permeability, leading to the carbon efflux from roots to rhizosphere. Thirdly, released carbon flux recruited beneficial rhizobacteria (e.g., strains LSS1 and LSS2) to help plant combat organic pollutant stress. Fourthly, root ROS stimulated endogenous NO production that facilitated the colonization of recruited rhizobacteria in plants. These results suggest that local exposure of pollutants triggers systemic acquired acclimation (SAA) across the entire plant with the assistance of rhizomicrobiota through long-distance ROS signaling.

The establishment of SAA requires rapid and long-distance signaling across plant tissues³³. Here, it was found that foliar exposure to organic pollutants not only induced local ROS burst in leaves, but also boosted ROS accumulation in other organs (e.g., stems and roots), suggesting that ROS may be a systemic signaling in this process. In response to stimuli such as light or wounding, plants exhibit systemic ROS accumulation within minutes³⁴. However, in this study, ROS waves in stems and roots also occurred but delayed, occurring 3-6 hours after foliar treatment with organic pollutants. Unlike the immediate cellular injury caused by light or wounding stress, plants may take longer to absorb and to initiate stress responses. Cadmium stress could induce a ROS burst in the tips of the exposed roots and trigger systemic ROS signaling to the unexposed roots³⁵. The Ca²⁺-RBOH and BrRBOHs pathways confirmed that plant systemic ROS wave in response to foliar pollutant exposure can be achieved through RBOH-dependent ROS production triggered by Ca²⁺ signals in various tissues, but not involving direct ROS diffusion^36,37. Interestingly, four RBOH genes (BrRBOH03, BrRBOH07, BrRBOH08, and BrRBOH11) in stems and four RBOH genes (BrRBOH03, BrRBOH06, BrRBOH12, and BrRBOH13) in roots were selectively upregulated by organic pollutants, suggesting their involvement in systemic ROS production²⁹. Overexpressing BrRBOH13 confers plant tolerance against heavy metal but not saline stress, suggesting the functional diversity of BrRBOH in regulating plant acclimation to different stressors²⁹. Although the functions of BrRBOH genes are not fully understood, studies on Arabidopsis homologs offer insights. For example, AtRBOHC and AtRBOHE, homologs of BrRBOH03, exhibit high ROS-producing activity regulated by Ca²⁺, implying that BrRBOH03 and BrRBOH06 may contribute to ROS production under pollutant stress^29,38,39.

The long-distance transmission of ROS signaling can be mediated through two primary mechanisms. In plants, the central receptor of ROS is H₂O₂-induced Ca²⁺ increases 1 (HPCA1), a leucine-rich-repeat receptor-like kinase⁴⁰. In Arabidopsis in response to local high light stress, AtRBOHD and AtRBOHF produce apoplastic ROS that can be sensed by HPCA1 in neighboring cells. Apoplastic ROS oxidizes the extracellular cysteine residues to activate HPCA1. Then HPCA1 simulates Ca²⁺ influx by phosphorylating Ca²⁺-permeable channel mechanosensitive ion channel-like 3 (MSL3). This elevated cytosolic Ca²⁺ triggers a kinase cascade involving a set of Ca²⁺-related proteins, further activating AtRBOHD and AtRBOHF to lead to apoplastic ROS production. This positive feedback loop ultimately facilitates systemic ROS wave to propagate from locally-stressed area throughout the entire plant⁴¹. In zebrafish larvae, long-distance ROS signaling occurs via an ROS gradient from wound margin to adjacent tissue⁴². In this process, Ca²⁺-dependent dual oxidase is required for ROS production, further leading to the rapid recruitment of leukocytes to wound sites. In the present study, both Ca²⁺-RBOH signaling and ROS gradient from leaves to roots were observed. Further research is needed to determine whether foliar pollutant exposure induces ROS signaling through similar mechanisms.

The cell membrane plays an essential role in substance exchange between roots and rhizosphere. ROS can cause lipid peroxidation by reacting with membrane lipids, leading to the loss of membrane integrity and increasing permeability⁴³. Therefore, the increase in membrane permeability will further facilitate the release of root exudates found. Additionally, it was also found that root exudates contained 18-carbon unsaturated fatty acids (e.g., linoleic acid) in response to foliar exposure of organic pollutants. These metabolites, key components of cell membrane lipids, are important for the maintenance of membrane fluidity and permeability⁴⁴. ROS regulates lipid biosynthesis and metabolism by triggering multiple signaling pathways, such as histidine kinases (HiKs)- and receptor-like protein kinases (RLKs)-sensing pathways, mitogen-activated protein kinase (MAPK) signaling cascade, and bZIP and MYB transcription factors⁴⁵. ROS directly regulates these functional proteins via cysteine thiol-based oxidative post-translational modifications, enabling redox signaling transduction⁴⁶. Although further research is needed to clarify ROS-mediated membrane lipid remodeling, the results of this study suggest that ROS increases root cell membrane permeability, promoting the release of root exudates in response to foliar pollutant exposure.

Environmental stress can disrupt plant metabolic homeostasis⁴⁷. Plant carbon flux is crucial for plant-environment interactions. Stress-induced carbon flux directs resources to stressed plant organs to support energy needs⁴⁸. Here, we found that foliar exposure to organic pollutants stimulated carbon efflux from roots, suggesting systemic carbon flux to support non-stressed organs during acclimation. Photosynthesis in leaves primarily produces organic carbons, which are then distributed throughout the whole plant via vascular system (e.g., xylem and phloem)⁴⁸. Although the role of ROS in modulating vascular carbon flux remains unclear, the findings of this work support ROS signaling role in promoting root carbon release in response to foliar exposure to organic pollutants.

Plants, in response to biotic and abiotic stress, can mediate the release of root exudates, such as organic acids, amino acids, and hormones, to recruit rhizosphere bacteria^49,50. Nonetheless, the upstream signaling pathways regulating the release of these exudates under stress remain elusive. A recent study demonstrated that local bacterial infection triggers RBOH-dependent ROS production that acts a mobile signaling, activating salicylic acid biosynthesis in distal parts of a plant for the establishment of systemic acquired resistance (SAR)⁵¹. In this study, it was found that a similar signaling response to local exposure to organic pollutants triggered long-distance movement of ROS signaling throughout the rest of the plant. Mobile ROS signaling could promote plant carbon efflux to recruit rhizosphere microbes in response to foliar pollutant exposure, suggesting that ROS acts to influence the rhizosphere environment rather than directly enhancing plant intrinsic tolerance. The increased carbon efflux could affect the rhizosphere microbial community in multiple ways. Primary metabolites such as amino acids and organic acids provide universal carbon sources for rhizosphere bacteria⁵². Additionally, certain secondary metabolites, including phenolic acids, flavonoids, and glucosinolates, not only exhibit antibacterial activity but also selectively recruit specific beneficial bacteria in the rhizosphere⁵³. We found that both Sphingomonas sp. LSS1 and Lysobacter sp. LSS2 displayed a strong ability to chemotax toward and utilize root exudates from stressed plants, leading to their enrichment in the rhizosphere. Furthermore, these strains exhibited distinct preferences on the root exudate metabolites, suggesting that they are more likely to coexist rather than compete in the rhizosphere. This supports the concept that ROS is a universal long-distance signal initiating SAA through diverse defense strategies, including plant-rhizobacteria interactions.

Besides of the systemic activation of plant intrinsic tolerance upon local abiotic stress or pathogen infection^51,54, we identified another strategy that mobile ROS signal could act as a messenger asking help from rhizobacteria through root exudates. These rhizobacteria assist plants to establish SAA together by promoting plant growth and organic pollutant degradation. In plant-pathogen interactions, the “cry-for-help” response has been associated with the capability of plant roots in recruiting rhizomicrobes to either antagonize pathogens or improve plant fitness in situ, which mainly occurs in soil-borne disease^55,56. Foliar activation of pattern-triggered immunity (PTI) has been shown to induce rhizomicrobial recruitment via root exudates, although the long-distance signals involved remain unknown⁵⁷. Having linked local stress sensing in the leaf to rhizomicrobial recruitment in the root, long-distance ROS signaling advances the understanding for plant “cry-for-help” responses in plant acclimation.

Recruited rhizomicrobes regulate plant physiology in numerous ways, but the colonization is an important step. NO, a multifunctional signaling molecule in plants, facilitates rhizobacterial colonization by acting downstream of ROS in roots, likely through root cell wall loosening. The interaction between plant endogenous NO and rhizomicrobes remains elusive. NO regulates cell wall components (e.g., pectin, cellulose, and hemicellulose)^58,59, in coordination with other signaling molecules such as gibberellic acid⁶⁰, auxin⁶¹, and Ca²⁺ influx⁶², etc. Our findings suggest that in organic pollutant-stressed plants, root endogenous NO could act downstream of ROS to enhance root cellulose activities, leading to loosening of the cell wall that facilitates the colonization of beneficial bacteria recruited by stressed plants. For plant tissues in direct contact with hazards, NO-strengthened cell walls may work as an important physical barrier to prevent pathogen invasion or pollutant entrance^60,63. Foliar pollutant exposure failed to trigger endogenous NO signaling directly in leaves. Instead, an NO response was found in roots, inducing root cell wall loosening. We cannot exclude the possibility that organic pollutants at higher concentrations trigger NO signaling in leaves. However, NO-loosened root cell walls help plant combat pollutant stress by acting as a second messenger to promote the colonization of beneficial rhizobacteria. This indicates the functional diversity of NO in plants facing different environmental scenarios.

In summary, we demonstrate that ROS functions as a long-distance signal linking foliar pollutant stress and rhizosphere modulation (Fig. 6). Foliar exposure to organic pollutants triggers long-distance ROS signaling that moves to roots and initiates two strategies involving rhizobacteria recruitment and colonization via root exudates and NO signaling, respectively. This study provides insights into how plants sense local stress to establish SAA with the assistance of rhizomicrobiota, emphasizing the role of ecological harmony in plant survival. Further studies will focus on the effects of phyllosphere microbes on plant resistance to organic pollutants, as well as their migration within plants.

**Fig. 6: Conceptual model for the function of ROS signaling as a long-distance signal linking foliar pollutant stress and rhizosphere modulation.**

Methods

Analysis of rhizosphere bacteria

Topsoil was collected from paddy fields at the Jiangsu Academy of Agricultural Sciences, Nanjing, Jiangsu Province, China, and was air-dried, passed through a 10-mesh sieve, and thoroughly mixed. The soil composition was as follows: 30% sand, 53% clay, and 17% silt, with an organic matter content of 2% and a pH of 6.1. B. rapa seedlings were planted in plastic pots with 110 g soil and were maintained in a growth chamber under the experimental conditions (light intensity of 200 μmol m⁻² s⁻¹, 72% humidity, a temperature of 30 °C, and a daily light-dark cycle of 10–14 h). After 20 days, every ten vegetable plants were leaf-sprayed by a 10 mL aqueous solution containing 0.1% polysorbate-80 and individual organic pollutants (Thia, Teb, Ace, Phen, and Trich; purity >99%, Sigma-Aldrich Chemical Co.) at a concentration of 10 mg/L for each organic pollutant, and the control group was leaf-sprayed by an equivalent aqueous solution containing 0.1% polysorbate-80. Before spraying, the tops of the pots were covered with plastic film to prevent organic pollutant contamination. Two weeks after treatment, rhizosphere soils from each group were collected for bacterial community analysis and isolation, as well as for the detection of pollutants and their intermediates.

Bacterial DNA was extracted from rhizosphere soil with an E.Z.N.A.® Soil DNA Kit (Omega Bio-tek, Norcross, GA, United States) and was analyzed using the MiSeq platform (Illumina, San Diego, United States) to sequence the V3-V4 region of the 16S rRNA gene. The operational taxonomic units (OTUs) were then clustered using UPARSE software, with a cutoff value of 97% similarity. Taxonomy assignment of OTUs was analyzed with the RDP classifier (http://rdp.cme.msu.edu/) against the Silva (SSU123) 16S rRNA database using a confidence threshold of 70%. Shannon diversity and PCoA based on Bray‒Curtis distances were calculated at the OTU level. Linear discriminant analysis effect size (LEfSe) with two-sided Wilcoxon rank-sum test was used to compare genera between the organic pollutants and control groups, with taxa considered significantly different only when P < 0.05 and LDA effect size > |2.5|.

Feedback effect of rhizosphere beneficial genera

Isolation of rhizosphere bacteria

The rhizosphere soil (0.5 g) from the Thia treatment described in Section 4.1 was mixed with a 20 mL sterilized phosphate-buffered saline (PBS) buffer at pH 7.0. Following incubating for 30 min at 37 °C, the supernatant was serially diluted and spread on Luria-Bertani (LB) agar plates. Individual colonies were isolated and identified using 16S rRNA gene sequencing⁶⁴. Bacteria from Sphingomonas and Lysobacter genera were successfully isolated, including representative strains, Sphingomonas sp. LSS1 and Lysobacter sp. LSS2. Both strains were domesticated by exposure to progressively higher concentrations of rifampicin. Strains LSS1 and LSS2 stably expressing the Red Fluorescent Protein (LSS1-rfp and LSS2-rfp, respectively) were obtained after plasmid pPROBE-pTetR-RFP infection using the triparental conjugation method¹⁶.

Degradation of Thia in vitro

Strains LSS1 and LSS2 were separately inoculated in a 50 ml LB medium and incubated at 150 rpm at 37 °C overnight. Cell pellets were rinsed three times with PBS (pH 7.0) and resuspended in PBS to an OD_600nm of 0.5. The suspension (1.0 mL) of individual LSS1 and LSS2, as well as their proportional synthetic microbial community, was then transferred to a 250 mL Erlenmeyer flask with a 100 mL sterilized mineral salt medium (MSM) solution and Thia (5 mg/L). The flask was kept in shaker at 150 rpm at 37 °C for 5 days and the solution was collected for Thia analysis.

Plant growth and Thia degradation in vivo

B. rapa seedlings were planted in pots with sterilized vermiculite and watered with half-strength Hoagland nutrient medium. After 20 days of cultivation, the root of plants was separately inoculated with 1.0 mL of LSS1 (OD_600nm = 0.5), LSS2 (OD_600nm = 0.5), and their proportional synthetic microbial community, and the control group was treated with 1.0 mL sterile water. Plant weights were recorded following 20 days of cultivation. Besides, two days after inoculation with bacteria, a portion of seedlings was leaf-sprayed with a 10 mL aqueous solution with 0.1% polysorbate-80 and Thia (10 mg/L), and the shoot of these plants was collected for Thia analysis during the subsequent 10 days.

Change of vegetable physiological metabolism and root exudates

Vegetable sample collection

Hydroponic B. rapa seedlings were cultivated in amber glass bottles with a 50 mL of half-strength Hoagland nutrient medium for a 20-day incubation period in a controlled environment chamber. Application of pesticides and organic pollutants followed the protocol outlined in Section 2.1.1. Additionally, vegetables were sprayed with 10 mg/L of Thia and 100 μM lanthanum chloride (Ca²⁺-permeable channel blocker). After 48 h application, plant shoots and roots were collected for analysis of physiological indexes and linoleic acid. Besides, six seedlings from each group were washed three times with sterile water and then transferred to amber glass vessels with a 60 mL sterilized water for an additional two-day incubation. After incubation, the remaining aqueous solution (root exudates) was filtered through a 0.4 µm filter membrane for analyses of metabolic profiling, bacterial culture, and organic pollutants.

Determination of plant physiological parameters and the expression of BrRBOH genes

The shoots of vegetables exposed to individual organic pollutants, as described in Section 4.3.1, were analyzed for physiological indexes. The contents of ROS, NO, and Ca²⁺ were determined by using commercial test kits S0033S, S0023, S1063S, respectively (Beyotime Biotechnology, Shanghai, China). ROS in root tips was visualized using nitro-blue tetrazolium staining⁶⁵. Nitric oxide synthase activity of was determined by using commercial test kit S0025 (Beyotime Biotechnology, Shanghai, China). Nitrate reductase activity of was determined by using commercial kit AKNM001M (BoxBio Technology, Beijing, China). Root membrane permeability was characterized by electrolyte leakage, which was determined by a Mettler Toledo MC-126 conductivity meter (Switzerland).

Quantitative Reverse Transcription-Polymerase Chain Reaction (qRT-PCR) was used to determine the relative expression of BrRBOH genes, including BrRBOH01 and BrRBOH03 ~ BrRBOH14, in plant tissues. The GeneJET Plant RNA Purification Kit (Thermo Fisher Scientific) was used to extract total RNA from plant roots. Reverse transcription was performed using SynScript^®Ⅲ RT SuperMix (Tsingke Biotechnology, Beijing, China) to obtain cDNA. Then the cDNA was used as template to perform PCR analysis using a QuantStudio stepone Plus (ABI, Thermo, United States) and the ArtiCanCEO SYBR qPCR Mix (Tsingke Biotechnology, Beijing, China) under the following conditions: initial denaturation at 95 °C for 5 min, followed by 40 cycles of 95 °C for 15 s, 60 °C for 20 s, and 72 °C for 20 s. The relative expression of each gene was normalized based on the abundance of internal standard gene BrActin. The primer sequences are listed in Supplementary Table 1.

Root exudate analysis

Root exudate solution mentioned in Section 4.3.1 was freeze-dried, weighed, redissolved in 2.5 mL of an aqueous solution of 80% methanol, vortexed 15 min, and centrifuged at 12,000 × g for 15 min¹⁷. The final supernatant was analyzed by LC-QTOF/MS (Shimadzu LC-20A HPLC coupled with a Sciex TripleTOF 5600 + ). Mass spectrometry was performed using electrospray in both positive and negative ionization modes. An XSelect HSS T3 column (4.6 × 150 mm, 3.5 µm; Waters Corporation) was used for chromatographic separation. The mobile phase A was 0.1% formic acid and 5 mM ammonium formate in water for positive and negative ionization, respectively, and the mobile phase B was HPLC-grade acetonitrile. The flow rate was set to 0.30 mL/min, and the gradient program for mobile phase B was as follows: 0−3.0 min, 1% B; 3.1−24.0 min, 1−100% B with linear ramp; 24.1−32.0 min, 100% B; 32.1−37.0 min, 1% B. The information about MS peaks, MS matrix, MS/MS spectra, and retention times was collected from the raw LC-QTOF/MS data using MS-DIAL software. Peak identification was performed by calculating the similarity of the retention time, mass accuracy, and MS/MS profile against publicly available databases, and confirmed with chemical standards where applicable. PCA was performed based on MS matrix. Metabolites with significant changes were confirmed by Student’s t test with a P-value < 0.05 and partial least squares discriminant analysis with a VIP value > 1.

Interactions of root exudates and bacterial strains

Bacterial preference for root exudates

Fifty milliliters of plant root exudate solution collected from the Thia treatment, as described in Section 4.3.1, were inoculated with either Strain LSS1 or LSS2 (OD_600nm = 0.1)⁶⁶. After a 6 h incubation at 37 °C, 30 mL of the solution was collected, filtered through a 0.44 μm membrane to remove bacteria, freeze-dried, redissolved, and analyzed by LC-QTOF/MS. The procedures of sample pretreatment and LC-QTOF/MS analysis are described in Section 4.3.3.

Chemotaxis analysis

A capillary method was used to test the chemotaxis of strains LSS1 and LSS2 towards root exudates and ROS from the Thia-treated and control groups⁶⁷. Ten microliters of root exudate solution or H₂O₂ (5 mM) was placed into a microcapillary tube and maintained in a 500 μL MSM solution containing either Strain LSS1 or LSS2 (OD_600nm = 0.1) for 30 min. The solution in the tube was diluted with PBS buffer at pH 7.0 and spread on LB plates to determine CFU counts after 24 h of incubation at 37 °C.

Exposure of plants to H₂O₂, SNP, and their inhibitors and scavengers

Hydroponic plants were separately treated with Thia, H₂O₂ (ROS donor), H₂O₂ + NAC (ROS scavenger), Thia + NAC, SNP (NO donor), SNP + c-PTIO (NO scavenger), and Thia + c-PTIO, respectively. For Thia treatment, plant seedlings were sprayed by 10 mg/L of Thia, as described in Section 4.1.1. Other reagents were added in half-strength Hoagland nutrient medium. The concentrations of H₂O₂, NAC, SNP, c-PTIO in the solutions were set to 100 mM, 2 mM, 500 μM, and 100 μM, respectively. After treatment for 24 hours, root samples were collected to measure the levels of ROS, NO, and cellulose activity. ROS and NO contents were determined using the kits mentioned in 4.3.2. Cellulose activity was determined using test kit BC2545 (Solarbio Life Sciences, Beijing, China).

Root cell wall integrity was evaluated using fluorescent probe Calcofluor white (Genemed Biotechnologies, Torrance, USA) fluorescence microscope (355 nm for excitation, 440 nm for emission). The remaining plants were transferred to a sterile water solution containing individual strains of LSS1-rfp or LSS2-rfp (OD600_nm = 0.1). Following 24 h of cultivation, plant roots were surface-sterilized and ground with 1 mL PBS (pH 7.0), and the solution was spread on LB agar plates containing rifampicin at the concentration of 300 mg/L. Bacterial colony-forming units were counted 2 days after incubation at 37 °C.

Root exudate release under ROS, NAC, and linoleic acid existing

Hydroponic plants were cultivated in half-strength Hoagland nutrient medium and were separately treated with H₂O₂, H₂O₂ + NAC, Thia + NAC, and linoleic acid (50 μg/L). After a duration of 24 h, plant tissues were collected to assess ROS levels, and the solutions of root exudates were collected for further metabolomics analysis, as described in Section 4.3.

Analysis of organic pollutants, their intermediates, and linoleic acid

A 0.50 g sample of plant tissues and rhizosphere soil or 0.5 mL of MSM solution was extracted with 4 mL of acetonitrile, followed by vortex mixing for 10 min and centrifugation at 3802 × g for 10 min. The clear supernatant was treated with a mixture containing 50 mg of graphitized carbon black and 150 mg magnesium sulfate. After vortexing for 2 min, the resulting solution (2 mL) was collected, and 1 g of sodium chloride was added. The mixture was vortexed for 10 min and centrifuged at 3802 × g for another 10 min. The clear supernatant was filtered through a 0.22 μm membrane. Thia, Teb, Ace, and linoleic acid were analyzed using LC-QQQ/MS (AB Sciex 5500+ triple quadrupole mass spectrometer) with a C18 reversed-phase column (ZORBAX SB-C18, 4.6 mm × 250 mm, 5 µm). Phen and Trich were analyzed using an Agilent 7890 gas chromatograph coupled with Agilent 7000D mass spectrometer (GC-MS) with a DB-5 capillary column (30 m × 0 25 mm × 0.25 μm). The limits of quantification (LOQs) of Thia, Teb, Ace, Phen, and Trich in plant tissues and rhizosphere soil were 1.1 μg/kg, 1.6 μg/kg, 1.3 μg/kg, 4.6 μg/kg, and 6.5 μg/kg, respectively. The LOQs of Thia in MSM solution and linoleic acid in plant roots were 0.9 μg/L and 1.8 μg/kg, respectively. Other information for analysis of organic pollutants and linoleic acid is shown in Supplementary Table 2-3.

Additionally, a 0.50 g sample of plant tissues and rhizosphere soil was extracted with 4 mL of an aqueous solution of 80% methanol, vortexed 15 min, and centrifuged at 12,000 × g for 15 min. The final supernatant was analyzed by LC-QTOF/MS, and the equipment conditions are described in Section 4.3.3. The MS-FINDER software was used to identify pollutant intermediates based on MS and MS/MS spectra⁶⁸.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The 16S rRNA gene sequencing data are available in the NCBI SRA under BioProject PRJNA1290846. The metabolomics raw data are available in the figshare database [https://doi.org/10.6084/m9.figshare.30000316.v1]. Source data are provided with this paper.

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Acknowledgements

This research was financially supported by the National Natural Science Foundation of China (Grant NO. 32272600 and NO. 32272583) and Fujian Provincial Science and Technology Plan Project (Grant No. 2022N0026). We thank Prof. Todd Alan Anderson from Texas Tech University for writing the manuscript.

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Authors and Affiliations

Jiangsu Key Laboratory for Food Quality and Safety-State Key Laboratory Cultivation Base, Ministry of Science and Technology, Nanjing, China
Yong Li, Kaiwei Zhang, Hui Zhang, Mei Li, Liya Ma, Yang Liu, Fayun Feng, Jing Ge, Jian Chen & Xiangyang Yu
Institute of Food Safety and Nutrition, Jiangsu Academy of Agricultural Sciences, Nanjing, China
Yong Li, Kaiwei Zhang, Hui Zhang, Mei Li, Liya Ma, Yang Liu, Fayun Feng, Jing Ge, Jian Chen & Xiangyang Yu
Department of Entomology, Michigan State University, East Lansing, MI, USA
Wayne Jiang
Jiangsu Key Laboratory for Bioresources of Saline Soils, School of Wetlands, Yancheng Teachers University, Yancheng, China
Xiangyang Yu

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Yong Li
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Kaiwei Zhang
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Hui Zhang
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Mei Li
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Liya Ma
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Yang Liu
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Fayun Feng
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Wayne Jiang
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Jing Ge
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Jian Chen
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Xiangyang Yu
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Contributions

X.Y., J.C. and Y.L. designed the study. Y.L., J.C. and K.Z. performed the experiments and analyzed the data. H.Z., M.L. and L.M. performed data deposition. Y.L., F.F. and J.G. contributed to data analysis and interpretation. Y.L. wrote the manuscript. X.Y., J.C. and W.J. edited the manuscript. All authors discussed the results and reviewed the manuscript.

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Correspondence to Jian Chen or Xiangyang Yu.

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Li, Y., Zhang, K., Zhang, H. et al. Organic pollutant-induced long-distance ROS signaling drives plant systemic acquired acclimation via rhizomicrobiota. Nat Commun 16, 9077 (2025). https://doi.org/10.1038/s41467-025-64138-y

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Received: 21 November 2024
Accepted: 05 September 2025
Published: 13 October 2025
DOI: https://doi.org/10.1038/s41467-025-64138-y