BRAHMA represses STOP1-NRT1.1 module to control plant rhizosphere alkalization and acid stress adaptation

Ye, Jia Yuan; Tian, Wen Hao; Zhang, De Rui; Yu, Qian Yi; Xu, Ji Ming; Ding, Wo Na; Li, Gui Xin; Yan, Jing Ying; Jin, Chong Wei; Benhamed, Moussa; Zheng, Shao Jian; Ding, Zhong Jie

doi:10.1038/s41467-026-69905-z

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Published: 23 February 2026

BRAHMA represses STOP1-NRT1.1 module to control plant rhizosphere alkalization and acid stress adaptation

Nature Communications volume 17, Article number: 3084 (2026) Cite this article

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Abstract

Covering 40–50% of world’s arable lands, acidic soils pose a major constraint on global crop productivity by severely restricting root development and nutrient acquisition. The Arabidopsis C2H2-type transcription factor STOP1 plays a fundamental role in mitigating acid stress by activating H⁺/NO₃⁻ symport via NRT1.1, thereby driving rhizosphere alkalinization to protect root growth and improving nitrogen use efficiency (NUE). However, the upstream regulation of this pH-responsive STOP1-NRT1.1 pathway remains poorly defined. Here, we identify the central SWI2/SNF2-type ATPase BRAHMA (BRM) as a key suppressor of the STOP1 pathway. BRM physically interacts with STOP1 and occupies the genomic region of NRT1.1, repressing the STOP1-dependent activation of NRT1.1 expression and consequently limiting NO₃⁻ uptake and rhizosphere alkalization under chronic acidity. Genetic epistasis analysis using brm stop1 and brm nrt1.1 double mutants establish BRM as an upstream regulator of this signaling module. Notably, low pH rapidly triggers BRM degradation independent of NO₃⁻ availability, thereby relieving its repression on the STOP1-NRT1.1 pathway. This dynamic BRM disintegration enables robust induction of H⁺-coupled NO₃⁻ uptake, remodeling the rhizosphere pH landscape to foster optimal root growth under acidity. Collectively, our findings uncover the BRM-STOP1-NRT1.1 axis as a central regulatory module integrating NO₃⁻ acquisition with pH homeostasis, offering a dual-benefit strategy for enhancing crop resilience to acid soils and reducing fertilizer-driven acidification through improved NUE.

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Introduction

Acidic soils occupy 40–50% of the world’s total arable lands, and their acidification pace is accelerating due to the widespread application of urea and ammonium-based fertilizers^1,2, posing a critical challenge to the sustainability of global agriculture. Acidic soils impose a vicious cycle: high proton (H⁺) concentrations in acidic soils severely limit crop productivity by inhibiting root growth and nutrient acquisition^3,4, while plants’ inability to efficiently utilize N fertilizers exacerbates soil acidification through residual H⁺ accumulation and also the risk of environmental pollution⁵. Molecular breeding of plants to improve nitrogen acquisition and adaptation to acidic soils could be a cost-effective solution to these problems⁶.

The C2H2-type transcription factor, SENSITIVE-TO-PROTON-RHIZO TOXICITY1 (STOP1), was identified as the first key regulator of plant H⁺ tolerance, with its disruption leading to over 70% inhibition of root growth under acidic conditions⁴. Subsequent lines of evidence have suggested that STOP1 has pleiotropic roles in plant response to various environmental cues, including ammonium toxicity, Pi deficiency, N deficiency, Ca²⁺ deficiency, salt stress, drought, and hypoxia stresses^{7,8,9,10,11,12,13,14,15,16}. Furthermore, STOP1 is functional conserved across land plant species^17,18. Among critical STOP1-regulated modules, our previous study demonstrated that the STOP1-NITRATE TRANSPORTER 1.1 (NRT1.1) module emerges as a critical buffer in response to low pH: STOP1 activates NRT1.1-mediated H⁺/NO₃⁻ symport, alkalizing the rhizosphere to protect roots and enhance N use efficiency⁵. However, the upstream mechanisms that underlie STOP1-enhanced plant adaptation to acidity remain unclear.

Soil acidification is a dynamically deteriorating process: even in moderately acidic soils (pH 5.0–5.5), repeated urea application can lower rhizosphere pH by 0.17–0.72 units over one growing season^1,19. If plant-induced H⁺ influx from root nutrient uptake could counteract (either fully or partially) H⁺ released during nitrification of ammonium-based fertilizers, this equilibrium could mitigate soil acidification, simultaneously benefiting agricultural ecosystem sustainability and crop productivity. In Arabidopsis, NRT1.1 serves as the major transporter for root NO₃⁻ uptake under adequate NO₃⁻ supplementation and acidic conditions^20,21. The NRT1.1-mediated H⁺/NO₃⁻ symport drives rhizosphere alkalization to protect roots from acid toxicity and enhance nitrogen use efficiency (NUE)⁵. Thus, adaptation to acidic stress requires plants to orchestrate both immediate stress responses and sustained transcriptional programming that stabilizes NRT1.1 expression to ensure sufficient NO₃⁻ uptake to neutralize protons derived from ammonium nitrification²². It’s worth noting that STOP1-dependent H⁺ tolerance is functionally restricted by its post-translational modifications and dosage sensitivity, thereby stringently restricting its capacity to activate NRT1.1 transcription. Furthermore, STOP1 overexpression not only produces growth penalties but also fails to confer chronic stress adaptation, while its upstream regulators governing transcriptional activity under H⁺ stress are largely unknown. Chromatin remodeling, by enabling sustained transcriptional memory during prolonged stress²³, emerges as a promising solution to circumvent these limitations through epigenetic stabilization of stress-responsive genes.

In this study, we identified BRAHMA (BRM), a switch defective/sucrose non-fermentable (SWI/SNF) ATPase, as a key repressor of the STOP1-mediated pathway. BRM directly occupies the genomic region of NRT1.1 and interacts with STOP1 to inhibit its trans-activation activity under acidity, thereby suppressing the STOP1-NRT1.1 module. Low pH rapidly triggers BRM degradation, relieving repression to augment H⁺-coupled nitrate influx and establish a rhizosphere pH optimum for root growth under acidic condition. Our findings may provide a strategy for simultaneously improving plant NUE to partially counter the H⁺ derived from nitrification of ammonium-based fertilizers and slowing soil acidification through the manipulation of the BRM-STOP1-NRT1.1 module.

Results

BRM physically interacts with STOP1

To identify potential regulators of STOP1 during H⁺ stress, we conducted a yeast two-hybrid (Y2H) screen using STOP1 as bait. Although pGBKT7-STOP1 exhibited weak autoactivation in the AH109 yeast strain, this background activity was effectively suppressed by 50 mM 3-amino-1,2,4-triazole (3-AT) (Supplementary Fig. 1). The screen yielded 75 positive colonies, corresponding to 23 putative STOP1-interacting proteins (Supplementary Data 1). Among these, BRAHMA (BRM), a core chromatin-remodeling subunit, was recovered in six independent clones, making it the most recurrent and therefore the highest-priority candidate among previously unreported STOP1 interactors. Region mapping assays showed that the N-terminal region of BRM (BRMn, amino acids 1 to 976) was both necessary and sufficient for their interaction (Fig. 1a–c). Since BRMn functions as a docking site for the recruitment of transcription factors²⁴, we employed this fragment, rather than full-length BRM, in subsequent biochemical assays. In Nicotiana benthamiana leaf BiFC experiments, co-expression of STOP1 and BRMn produced a robust nuclear fluorescence signal, confirming their interaction in planta (Fig. 1d). Furthermore, co-immunoprecipitation assay using Arabidopsis plants co-expressing pBRM:BRM-FLAG and pSTOP1:GFP-STOP1 (GFP-STOP1/BRM-FLAG) confirmed that GFP-STOP1 was specifically co-immunoprecipitated with BRM-FLAG after a 6-h low-pH and MG132 treatment (Fig. 1e). These results firmly establish that BRM interacts with STOP1in the nucleus.

Lack of BRM enhances plant growth and NO₃ ⁻ acquisition under H⁺ stress

To investigate the role of BRM in response to acidic stress, we performed a phenotypic analysis of two hypomorphic alleles of BRM (brm-3 and brm-20) and a complementation line (pBRM:BRM-FLAG/brm-3, hereafter referred to as COM). Under low-pH conditions, loss of BRM function significantly enhanced root growth, with root elongation in brm-3 and brm-20 mutants increasing by approximately 60% compared with that of wild-type (Col-0) plants, while the complementation of BRM fully restored the root growth of brm-3 mutants (Fig. 2a, b). Even after prolonged exposure, brm-3 roots remained longer than those of Col-0 under acidic pH, whereas under neutral pH the brm-3 mutant displayed significantly shorter root elongation compared with Col-0 (Supplementary Fig. 2). These results indicate that BRM negatively regulates root tolerance to acidic stress.

**Fig. 2: Downregulation of BRM enhances plant tolerance to low pH and stimulates nitrate uptake under acidic conditions.**

Given that BRM degradation is crucial for maintaining root apical meristem integrity by alleviating DNA double-stand breaks (DSBs) induced under high-B conditions²⁵, we examined whether the enhanced H⁺ tolerance of brm-3 mutants is related to altered DSBs levels. However, the expression of DSBs-induced genes (GR1, RAD51, and BRCA1) was neither significantly induced by low-pH treatment nor affected by BRM or STOP1 activity (Supplementary Fig. 3). Consistently, 1-day low-pH treatment did not significantly alter WT root DSBs level, suggesting that plants tolerance to acidic stress is not closely related with DSBs. Furthermore, the DSBs level of the acid-tolerant brm-3 mutant showed similar level as that of the acid-sensitive stop1 mutant, though they were slightly suppressed when compared to Col-0 (Supplementary Fig. 4). These results indicate that BRM-regulated H⁺ tolerance is not primarily associated with its role in DNA damage repair. Given that STOP1-improved H⁺ tolerance and NUE are closely linked to enhanced root NO₃⁻ uptake⁵ and that BRM physically interacts with STOP1 (Fig. 1), we speculated that BRM may regulates H⁺ tolerance in a NO₃⁻-depended manner. To test this notion, we compared the effects of different concentrations of NO₃⁻ supplies (0.06 and 6 mM) on root elongation of Col-0, brm-3, and stop1 plants, with 1 mM NH₄⁺ supplied as an additional nitrogen source. Under low-pH treatments with 0.06 mM NO₃⁻, root growth of Col-0, brm-3, and stop1 mutants was comparable (Fig. 2c, d). In contrast, brm-3 mutants showed significantly enhanced root growth compared to Col-0 under low-pH treatments with 6 mM NO₃⁻, while stop1 mutants exhibited reduced root elongation (Fig. 2c, d). These results suggest that BRM-regulated H⁺ tolerance depends on NO₃⁻ availability.

Notably, both brm-3 and brm-20 are weak alleles. While brm-3 harbors a T-DNA insertion that disrupts the bromodomain in the C terminus, brm-20 is predicted to result in a truncation of a small segment at the C terminus. These mutations do not appear to be essential for chromatin remodeling activity, but may affect BRM stability or its tethering to chromatin^26,27,28. To confirm the specific role of BRM in acid stress response, we further analyzed the phenotype of BRM-null mutant (brm-1), which is sterile and exhibits severe developmental defects^27,29,30. As shown in Fig. 2e, f, root elongation in the brm-1 mutant was largely insensitive to low-pH stress, in stark contrast to the significant inhibition observed in Col-0 plants. Moreover, decreasing the concentration of NO₃⁻ from 6 mM to 0.06 mM significantly inhibited root growth of Col-0 but barely affected that of brm-1 mutants under low-pH treatments, demonstrating that BRM-repressed H⁺ tolerance depends on NO₃⁻ availability.

Since NRT1.1 mediates the majority of root NO₃⁻ uptake under low-pH condition and the low pH-induced transcription of NRT1.1 is controlled by STOP1^5,21, we analyzed the expression of NRT1.1 in the roots of Col-0, brm-3, and stop1 mutants under different pH conditions with 6 mM NO₃⁻ supplementation. As expected, low pH-induced NRT1.1 expression was remarkably enhanced in roots of brm-3 and brm-1 mutants compared with that in Col-0 plants (Fig. 2g, h). To further investigate whether BRM and NRT1.1 function in the same pathway in response to H⁺ stress, we generated a brm nrt1.1 double mutant by crossing the nrt1.1-1 and brm-20 mutants (Supplementary Fig. 5). We were unable to obtain brm-3 nrt1.1 double mutant due to the severely reduced fertility of brm-3, as recently reported³¹. In low-pH agar medium, brm nrt1.1 plants exhibited root growth comparable to that of the nrt1.1-1 single mutants (Fig. 2i, j), suggesting that NRT1.1 is epistatic to BRM in the regulation of H⁺ tolerance. Collectively, these results indicate that BRM inhibits H⁺ tolerance by repressing the low-pH-induced expression of NRT1.1.

Lack of BRM enhances STOP1-activated NRT1.1 expression and NRT1.1 chromatin accessibility

To elucidate how BRM regulates NRT1.1 expression in response to H⁺ stress, we first assessed whether BRM affects STOP1 protein accumulation, as the induced NRT1.1 expression by low pH is known to be STOP1-dependent⁵. Under low pH condition, comparative analysis of GFP-STOP1 fluorescence in Col-0 versus brm-20 mutants (GFP-STOP1/brm-20) revealed no significant difference in STOP1 protein levels (Supplementary Fig. 6), suggesting that BRM does not repressed NRT1.1 expression by altering STOP1 protein accumulation. We next examined whether BRM modulates STOP1-dependent transcriptional activation of NRT1.1. Using a transient expression system in N. benthamiana leaves, we co-expressed STOP1 and BRM along with luciferase gene (LUC) reporter constructs driven by either the NRT1.1 promoter region (P1) or the 4^th intron (P2), both containing STOP1-binding sites^5,14 (Fig. 3a). In addition, NLP7, another well-known transcription factor that activates NRT1.1 transcription³², and the NRT1.1 promoter fragment (P3), which contains the binding regions of NLP7 but not STOP1 or BRM, were used as negative controls. As shown in Fig. 3b, c, BRM co-expression substantially decreased the STOP1-activated P1- and P2-derived LUC signals to an almost undetectable levels, suggesting that BRM indeed affects the activation of STOP1 on NRT1.1 expression. However, BRM co-expression had no significant impact on either the NLP7-activated P3-drived LUC activity driven by the P3 fragment or EV-resulted P3-mediated LUC activity (Supplementary Fig. 7). These results suggest that BRM specifically targets the STOP1-dependent activation of NRT1.1, rather than exerting a non-specific inhibitory effect on LUC expression. To assess whether this regulatory interaction occur in vivo, we carried out chromatin immunoprecipitation (ChIP) experiments. STOP1 enrichment at endogenous NRT1.1 chromatin regions was notably higher in brm-3 mutants than in Col-0 plants (Fig. 3d), confirming that BRM inhibits the binding of STOP1 to NRT1.1 genomic regions.

Fig. 3: BRM and STOP1 antagonistically regulate NRT1.1 expression. — **Fig. 3: BRM and STOP1 antagonistically regulate *NRT1.1* expression.**

To further investigate BRM’s autonomous capacity to inhibit NRT1.1 expression, we analyzed publicly available datasets using WashU Epigenome Browser (https://epigenomegateway.wustl.edu/) and Plant Regulomics (https://plantregulomics.org/). This analysis revealed that BRM is associated with NRT1.1 genomic regions harboring STOP1-binding motifs (Fig. 3e; Supplementary Fig. 8). This association was further confirmed by ChIP-qPCR, which showed BRM enrichment at these sites (Fig. 3f). Together, these findings support the idea that BRM antagonistically regulates NRT1.1 expression by interfering with STOP1 activity. Given that BRM modulates gene expression by altering chromatin accessibility^33,34,35, we next assess the chromatin state of the NRT1.1 locus using FAIRE-qPCR. Consistent with the ChIP-qPCR results, chromatin accessibility at the P1-1 and P2-1 regions of NRT1.1 was significantly increased in brm mutants compared to Col-0, with a stronger effect observed at P2-1 (Fig. 3g). These results indicate that BRM represses NRT1.1 expression by limiting chromatin accessibility at STOP1-binding regions, thereby reducing STOP1 occupancy and inhibiting NRT1.1 activation.

In addition, previous studies have shown that BRM interacts with the histone deacetylase HDA6 to repress transposons under various stresses by modulating DNA methylation and H3 acetylation levels^26,36. Our data also implicate HDA6 in this regulatory circuit: loss of HDA6 function enhanced H⁺ tolerance and increased NRT1.1 expression in root tips—the zones most sensitive to acidic stress and the key sites where STOP1 acts to prevent H⁺ toxicity⁵ (Fig. 3h–j). These findings suggest that BRM and HDA6 may cooperatively repress NRT1.1 through chromatin remodeling mechanisms that likely involve H3 acetylation and DNA methylation.

BRM acts genetically upstream of STOP1 in regulation of NO₃ ⁻ uptake and H⁺ tolerance

To clarify the genetic relationship between BRM and STOP1 in response to H⁺ stress, we generated a brm stop1 double mutant by crossing the brm-20 and stop1 mutants (Supplementary Fig. 9). When grown on low-pH agar medium, the brm stop1 double mutant displayed root growth similar to the stop1 mutant but notably shorter than that of Col-0 and brm mutant (Fig. 4a, b). In accord with the elevated NRT1.1 expression observed in roots of brm mutant, the brm plants also showed increased root NO₃⁻ uptake compared to Col-0 (Fig. 4c, d). However, both the enhancement of NRT1.1 expression and root NO₃⁻ uptake rate was completely abolished in the brm stop1 double mutant (Fig. 4c, d), indicating that the increased NRT1.1 expression and NO₃⁻ uptake in brm mutant requires functional STOP1. These results suggest that BRM exerts genetically upstream of STOP1 in regulating NO₃⁻ uptake and H⁺ tolerance under low pH conditions.

**Fig. 4: STOP1 functions genetically downstream of BRM in response to low pH.**

Lack of BRM enhances rhizosphere alkalization in response to H⁺ stress

Given the central role of the STOP1-NRT1.1 module in mediating H⁺-coupled NO₃⁻ uptake during acid stress, we hypothesized that targeted manipulation of BRM could promote a more favorable rhizosphere pH for root growth under low pH and curb N fertilizer-induced soil acidification. To test this, we first examined whether co-planting brm−3 and stop1 mutants could influence the root growth of stop1 mutants in low-pH agar media. Remarkably, when brm-3 and stop1 mutants were co-planted, stop1 root growth recovered to ~80% of brm-3 levels, versus only 57% in monoculture (Fig. 5a, b; Supplementary Fig. 10). Parallel experiments with Col-0 and brm-3 at pH 4.5 (accounting for inherent STOP1-NRT1.1 alkalinization capacity) similarly demonstrated improved Col-0 root growth in co-culture (Supplementary Fig. 10). These results suggest that loss of BRM function further promotes rhizosphere alkalinization, thereby enhancing root growth in response to acidic stress.

Fig. 5: Disruption of BRM enhances rhizosphere alkalization and H+ tolerance under acidic stress. — **Fig. 5: Disruption of BRM enhances rhizosphere alkalization and H⁺ tolerance under acidic stress.**

To further assess the role of BRM in regulating rhizosphere pH, seedlings were grown in acidic agar medium with or without 0.75 mM Homo-PIPES, a pH buffer used to minimize pH fluctuations. In the buffered acidic media, brm-3 mutants lost their growth advantage, exhibiting root elongation comparable to that of Col-0 and stop1 (Fig. 5c, d). Conversely, in unbuffered acidic media, brm-3 mutants showed dramatically improved root growth and enhanced rhizosphere alkalinization, as quantified by bromocresol purple staining and direct pH measurement after 6 h (Fig. 5e, f). In accordance with these results, the brm-1 mutant displayed significantly enhanced and rapid rhizosphere alkalinization compared to Col-0 (Supplementary Fig. 11). Taken together, these results indicate that loss of BRM enhances rhizosphere pH, thereby increasing H⁺ tolerance and offering a promising strategy to mitigate soil acidification under acidic conditions.

H⁺ stress induces BRM degradation

We next investigated the regulatory dynamics of BRM in roots under H⁺ stress. While low-pH treatment did not significantly alter the transcript levels of BRM (Fig. 6a), it substantially reduced the nuclear accumulation of BRM-GFP in root epidermal cells of pBRM:BRM-GFP/brm-20 transgenic plants (Fig. 6b). This decrease occurred independently of NO₃⁻ availability (Supplementary Fig. 12), and was detected as early as 2 h after low-pH exposure, preceding the induction of GFP-STOP1, which became evident at 4 h (Fig. 6b, c). This temporal hierarchy establishes BRM destabilization as an early event in the acid stress response, positioning BRM upstream of STOP1 signaling. Furthermore, the low pH-repressed BRM accumulation was restored by the treatment with the 26S proteasome inhibitor MG132 (Fig. 6d–f), suggesting that H⁺ stress likely promotes BRM degradation via the proteasome-dependent proteolysis. To investigate whether this degradation requires SnRK2 kinases, which are known to phosphorylate and inactivate BRM upon ABA treatment²⁴, we assessed the growth of snrk2.2/2.3 and snrk2.6 mutants. However, these mutants exhibited root growth comparable to Col-0 under both neutral and low-pH conditions (Supplementary Fig. 13), ruling out SnRK2s participation in BRM-mediated H⁺ tolerance. Overall, these findings support a model in which low pH triggers proteasomal degradation of BRM, thereby enabling activation of the STOP1-NRT1.1 module to enhance H⁺ tolerance.

**Fig. 6: Low pH induces BRM degradation.**

Discussion

Understanding how plants adapt to H⁺ stress and acquire nutrients under acidic conditions is critical for improving nutrient use efficiency and crop production in acidic soils. The H⁺-coupled NO₃⁻ uptake by the root, mediated by the STOP1-NRT1.1 module, is a key biological process for improving plant H⁺ tolerance^5,21,37,38. Here, we identified BRM as a pH-dependent repressor of STOP1-NRT1.1 module, revealing a vital regulatory layer in plant H⁺ adaptation (Fig. 6g). In this model, BRM physically interacts with STOP1 and binds directly to regulatory regions of NRT1.1, suppressing the STOP1’s transcriptional activation on NRT1.1. Low pH alleviates this repression by triggering BRM degradation, and thereby enables STOP1 to stimulate the H⁺-coupled NO₃⁻ uptake by NRT1.1, consequently enhancing rhizosphere alkalization, and establishing a favorable pH environment for root growth.

Despite the identification of STOP1’s role in H⁺ tolerance over a decade ago, the upstream regulatory mechanisms governing its activation and STOP1-improved H⁺ tolerance have long been enigmatic. Lines of evidences reveal that STOP1 is post-translationally modified in response to multiple stresses^{4,13,16,39,40,41,42}. Unlike the Al³⁺-induced post-translational modification of STOP1(e.g., deSUMOylation/SUMOylation, phosphorylation, and ubiquitination^{39,40,42,43,44,45,46}, our findings demonstrate that BRM physically interacts and suppresses STOP1 activity without affecting the STOP1 protein level, acting instead at the level of chromatin and transcriptional regulation (Fig. 3; Supplementary Fig. 3). Our identification of a BRM–STOP1 interaction is further supported by an independent high-throughput Y2H screen in which BRM was used as bait and STOP1 emerged as one of the strongest interactors⁴⁷. Functionally, BRM specifically antagonizes STOP1-dependent activation of NRT1.1 (Fig. 3; Supplementary Fig. 7). Genetic evidence reinforces this relationship: the brm stop1 double mutant phenocopied stop1 in root growth, nitrate uptake, and NRT1.1 expression (Fig. 4), and was clearly distinct from the WT and brm single mutant. This non-additive genetic behavior demonstrates that STOP1 is epistatic to BRM in controlling H⁺ tolerance and NRT1.1 expression. BRM establishes a low-activity baseline for the STOP1–NRT1.1 system, preventing unwarranted rhizosphere alkalinization and nitrate uptake under non-stress conditions. Crucially, exposure to low pH triggers rapid BRM degradation (Fig. 6), creating a temporal activation window in which STOP1 and NRT1.1 can be robustly induced only under bona fide acidic stress. In this way, BRM acts as a pH-responsive gatekeeper that ensures both stability and sensitivity of the STOP1 regulatory network and dynamically coordinates pH homeostasis with nitrate acquisition. This pH-sensitive switch enables rapid metabolic and rhizosphere remodeling in response to fluctuating acidity—an essential strategy for soil adaptation. Future studies should focus on how plants sense acidic stress signals and elucidating the molecular cues that trigger BRM degradation in response to acidic stress. Possible upstream regulators include pH-sensing kinases or ion channels that could link environmental signals to BRM stability and functions. In parallel, the mechanisms underlying nucleus-localized STOP1 enrichment in response to H⁺ stress remain to be elucidated.

Beyond NRT1.1, BRM–STOP1 cooperativity extends to multiple STOP1 targets. Two canonical STOP1-regulated genes, ALMT1 and CIPK23, were significantly upregulated in brm mutants and this induction was fully abolished by STOP1 disruption (Supplementary Fig. 14). Phenotypically, brm mutants displayed enhanced Al tolerance and increased sensitivity to phosphate deficiency—both STOP1-regulated processes—while brm stop1 double mutants again resembled stop1 (Supplementary Figs. 15 and 16). These findings indicate that BRM acts as a broad-acting negative regulator of STOP1-mediated stress responses, supporting a direct mechanistic role of BRM–STOP1 interaction rather than a locus-specific effect at NRT1.1 alone. Moreover, while STOP1 is indispensable for multiple stress responses, sustained high STOP1 activity is detrimental under normal conditions (Supplementary Fig. 17). By physically interacting with STOP1 to dampen its transcriptional activity and by occupying chromatin regions of STOP1 target genes to restrict accessibility, BRM constrains both the intensity and spatial range of STOP1 function. This repression is essential to prevent inappropriate activation of STOP1-mediated pathways and the associated growth penalties. We therefore propose that the BRM–STOP1 module may operate as a generalized regulatory logic for managing trade-offs between growth and stress resilience. BRM-mediated repression keeps STOP1 activity at an optimal low level under non-stress or mild stress conditions, preserving normal development. When stress intensifies, BRM inhibition is relieved, enabling precise and timely activation of STOP1-mediated adaptation programs. This dynamic repression–activation mechanism likely underpins plant acclimation to diverse and rapidly changing environmental stresses.

BRM, a core SWI/SNF chromatin-remodeling ATPase, has been implicated in a variety of stress responses, such as drought, phosphate deficiency, and gibberellin acid (GA) signaling^{24,25,33,36,47,48,49,50,51}. Our results revealed BRM directly represses NRT1.1 expression by modulating chromatin accessibility at its regulator loci and through physical interaction with its key activator STOP1. Importantly, this repression is relieved in brm mutants, which exhibit increased NO₃⁻ uptake and H⁺ tolerance under both neutral- and acidic conditions (Figs. 1 and 3). It’s worth noting that significant phenotypic divergence among brm alleles have been observed in response to low-pH treatment, which may due to the different functional domains affected among brm alleles. Using Y2H and BiFC assays, we showed that the N-terminus of BRM (BRMn, amino acids 1-976) is both necessary and sufficient for its physical interaction with STOP1 (Fig. 1). Notably, both brm-3 (harbors a T-DNA insertion disrupting the bromodomain in the C-terminus) and brm-20 (undergoes a small C-terminal truncation) mutants retain the intact N-terminal domain, thus maintaining the ability to interact with STOP1, which accounts for the partial retention of BRM’s repressive function on the STOP1-NRT1.1 module under low pH conditions. In contrast, the brm-1 null mutant, which lacks the entire protein including the N-terminal STOP1-interaction domain, exhibits a complete loss of BRM-mediated repression, leading to the full acid-tolerant phenotype. This underscores that the BRM N-terminus serves as the core domain for repressing the STOP1-NRT1.1 module. Furthermore, previous studies have showed that deletion of the C-terminal bromodomain (disrupted in brm-3) and the small C-terminal segment (truncated in brm-20) seems not to be essential for chromatin remodeling activity, but may affect BRM stability or its tethering to chromatin^26,27,28. Therefore, although these regions are not required for BRM-STOP1 interaction (Fig. 1), their disruption does affect acid stress tolerance (Fig. 2). These results, together with the finding that low pH rapidly promotes BRM degradation (Fi.6), suggest that BRM C-terminus also play roles in modulating acid stress adaptation, likely via fine-tuning BRM stability. Further studies are needed to elucidate the function and mechanisms of these functional domains in acid stress tolerance. In addition, several studies showed that BRM interacts with histone deacetylase HDA6 to repress transposons in response to several stresses through modulation of DNA methylation and histone H3 acetylation levels^26,36. Our data also point to a potential role for histone deacetylase HDA6 in cooperatively repressing NRT1.1 via chromatin remodeling mechanisms involving H3 acetylation and DNA methylation. Nevertheless, how BRM is recruited to the NRT1.1 loci and whether additional histone modifiers participate in this regulatory module require further investigation.

In agricultural contexts, excessive nitrogen fertilizers application, particularly urea-N fertilizer, has not only reduced NUE in crops but also significantly induced severe soil acidification^1,52,53. This acidification further slows down microbe-driven nitrification processes⁵⁴, prolonging the retention of fertilizer nitrogen in NH₃ form and increasing NH₃ volatilization potential. These sequential processes create a vicious cycle where continued acidification suppresses nitrification, while accumulated ammonium becomes increasingly prone to gaseous loss. Consequently, developing effective strategies for timely removal of H⁺ ions generated during nitrification represents a critical intervention point for improving NUE in agricultural systems. The BRM-STOP1-NRT1.1 pathway offers dual agronomic and ecological benefits by improving NUE and mitigating soil acidification driven by nitrogen fertilizers via enhancing NO₃⁻ uptake and rhizosphere alkalinization (Figs. 4 and 5). Theoretically, genetic manipulation of BRM to decouple its association with STOP1 and the genomic loci of NRT1.1 could optimize crop resilience in acidic soils while reducing environmental costs. Elucidating the evolutionary conservation of this pathway in staple crops, like rice or wheat, could unlock strategies for sustainable agriculture in acid-affected regions.

In summary, we identify BRM as a key regulator of the STOP1-NRT1.1 module, orchestrating rhizosphere pH homeostasis and nutrient acquisition under acidic conditions through modulation of H⁺-coupled NO₃⁻ uptake. This work bridges chromatin remodeling, H⁺ stress signaling, and nutrient acquisition, and provides a foundation for engineering crops with enhanced H⁺ tolerance and NUE. Further exploration of BRM’s degradation mechanism and broader its interaction network will further advance the development of resilient crop systems adapted to H⁺ stress and help address global soil acidification challenges.

Methods

Plant material

The Arabidopsis thaliana plants used in this work were all from Columbian (Col-0) background. The various mutants were stop1^KO (SLAK_114108), nrt1.1-1 (SALK_097431), tfl2-1 (CS3796⁵⁵), axe1-5³⁶, brm-1 (chr2-1)⁵⁶. The brm-3 (SALK_088462), brm-20 (SALK_002500), pBRM:BRM-GFP/brm-20, and pBRM:BRM-FLAG/brm-3 transgenic line used in this study has been previously described²⁶. The brm stop1 and brm nrt1.1 double mutants were generated by crossing the brm-20 and stop1 mutants or nrt1.1-1 mutants, respectively, and homozygous lines were identified by PCR. The pSTOP1:GFP-STOP1/pBRM:BRM-FLAG (GFP-STOP1/BRM-FLAG) line was generated by crossing the pSTOP1:GFP-STOP1/stop1(#B10) transgenic line⁸ with pBRM:BRM-FLAG/brm-3, and the homozygous lines were selected in agar media containing basta and hygromycin. The primers used are shown in Supplementary Data 2.

Plant growth condition

Two days of stratification were conducted at 4 °C before sowing the seeds. Surface-sterilized Arabidopsis seeds were sown on basal agar medium containing KNO₃ (6 mM), (NH₄)₂SO₄ (1 mM), NaH₂PO₄ (1 mM), MgSO₄ (500 μM), CaCl₂ (1 mM), H₃BO₃ (10 μM), MnSO₄ (0.5 μM), ZnSO₄ (0.5 μM), CuSO₄ (0.1 μM), (NH₄)₆Mo₇O₂₄ (0.1 μM), Fe-EDTA (50 μM), 1% agar (Sigma-Aldrich, A1296), 1% sucrose at pH 6.5 in a plant growth chamber (BPC600H, Fujian Jiupo Biotechnology, Co., Ltd.) at 22–23 °C under a 16-h light/8-h dark photoperiod cycle. Then, the 3-day-old plants were used for phenotype analysis in the above basal agar medium with an initial pH of either 4.7 or 6.5, and 0.75 mM Homo-PIPES were used for the buffering assay, as indicated in the figure legends. To analyze the effect of NO₃⁻ level on root growth in response to low pH, the media were supplemented with 0.06 or 6 mM KNO₃, while the concentration of (NH₄)₂SO₄ was reduced to 0.5 mM to avoid severe ammonium toxicity and the resulting differences in K concentrations were balanced by adjusting the K₂SO₄ concentration. To analyze the root growth of seedlings in response to low Pi, the Pi concentration in the media were reduced to 0.01 mM (low Pi). To analyze the root growth of seedlings in response to Al toxicity, seeds were directly sown on the control or Al treatment agar plates, prepared as previously described⁵⁷.

Yeast two-hybrid screen and assays

For screening, the full CDS of STOP1 was amplified and inserted into pGBKT7 using the EcorI and BamHI restriction sites resulting in a pGBKT7-STOP1 constructs. This gene construct was transformed into yeast AH109 cells. Transformants were selected on SD/-Trp medium (Takara), after which the autoactivation activity of the bait vector pGBKT7-STOP1 was tested by plating on SD/-Trp-His-Ade medium (Takara) supplemented with different concentrations of 3-amino-1,2,4-triazole (3-AT). Then, the pGBKT7-STOP1 bait vector was cotransformed with an Arabidopsis cDNA library into AH109 yeast competent cells according to the manufacturer’s instructions (Clontech). Transformed cells were plated on SD/-Trp-Leu-His-Ade medium supplemented with 50 mM 3-AT, following incubation on 30 °C. Plasmids were isolated from positive yeast clones for analysis of insert and sequencing.

For targeted Y2H assays, the coding region of BRM and truncated version of BRM was amplified and inserted into pGBKT7 using the EcorI and BamHI restriction sites. The full CDS of STOP1 was amplified and inserted into pGADT7 using the EcorI and BamHI restriction sites. Yeast AH109 cells were co-transformed with specific bait and prey constructs. Yeast growth was shown on SD/-Trp/-Leu and SD/-Trp/-Leu/-His/-Ade medium, following incubation on 30 °C. The primers used for generating the various clones are listed in Supplementary Data 2.

Bimolecular fluorescence complementation (BiFC) assays

The full CDS of STOP1 and coding region of truncated BRMn was individually inserted into p2YN and p2YC vectors (46) to fuse with the N-terminal or C-terminal fragment of YFP. The primers used to generate the various clones are listed in Supplementary Data 2. The different combinations of recombinant plastmids were co-infiltrated into young Nicotinana benthamiana leaves for 48 h. The YFP fluorescence was detected using a confocal laser-scanning microscope (LSM880; Zeiss) and indicated in green color.

Co-IP assay

To better reflect the authentic in vivo interaction status between BRM and STOP1, we specifically used the double-tagged genetic material (BRM-FLAG/GFP-STOP1) for Co-IP assays. However, under neutral-pH conditions, the abundance of GFP-STOP1 protein in this transgenic line is extremely low, making it technically challenging to capture the GFP-STOP1 protein through immunoprecipitation. To address this issue, 10-d-old pSTOP1:GFP-STOP1 and pSTOP1:GFP-STOP1/pBRM:BRM-FLAG with 6-h low-pH treatment to induce GFP-STOP1 protein accumulation and with MG132 to inhibit the degradation of BRM-FLAG protein. Then, seedlings were harvested for total protein extraction in RIPA lysis buffer (FD011; Fdbio Science Biotech Co., Ltd). After centrifugating at 13,000 × g for 10 min, 50 µL of supernatants were kept as input controls, whilst the rest was incubated with 30 µL of anti-FLAG magnetic beads for 2 h. After the beads were washed five times with TBS buffer, the proteins were eluted by adding 50 µL 2× SDS sampling buffer at 95 °C for 8 min. Then the eluted proteins and input controls were separated on 7.5% sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE). The protein of interest was detected with α-GFP (1:10000; ABclonal) and α-FLAG (1:10000; ABclonal) antibodies.

Gene expression assay

Total RNA was extracted from plant roots using the FreeZol Reagent (R711-01; Vazyme Biotech Co., Ltd, Nanjing, PRC), and the first-strand cDNA was synthesized using ReverTra Ace qPCR RT Master Mix with gDNA Remover (TOYOBO, Osaka, Japan). The expression of the corresponding genes was determined using the SYBR Green Realtime PCR Master Mix (TOYOBO). The gene-specific primers used for quantitative PCR were listed in Supplementary Data 2. Expression levels of each mRNA were determined and normalized with the expression level of UBIQUITIN 10.

Analysis of green fluorescent protein (GFP)

After different time of low-pH treatment, as indicated in figure legends, BRM-GFP and GFP-STOP1 expression in the roots of pBRM:BRM-GFP/brm-20 and pSTOP1:GFP-STOP1 transgenic plants was detected using a confocal laser-scanning microscope (LSM880; Zeiss). The excitation and emission wavelengths were 488 nm/500–530 nm for GFP protein. Images were analyzed using the ZEN 2012 Blue Edition and ImageJ2X software.

Analysis of pH and visualization of rhizosphere alkalization

The 10-day-old Arabidopsis seedlings were pre-treated at pH 4.7 agar medium for 2 days before bromocresol purple staining and the pH value measurement. To visualize rhizosphere acidification, roots were transferred to a basal agar media with an initial pH of 4.9 and pH indicator bromocresol purple (0.008%, Sigma-Aldrich). The concentration of NaH₂PO₄ was reduced to 0.1 mM in the used agar media. Images were acquired 12 h after staining. The pH of the agar rooting medium was measured as previously described⁵⁸. Briefly, the rooting medium was collected and frozen at –20°C for 4 h before thawed to free the aqueous phase from the agar. Then, the mixture was filtered at room temperature and the pH of the supernant was measured immediately using a pH electrode.

Chromatin immunoprecipitation (ChIP)-qPCR analysis

To assess the effect of BRM disruption on the binding ability of STOP1 to NRT1.1, protoplasts were isolated from the rosette leaves of 3-week-old WT or brm-3 plants, as described previously⁵⁹. The effector plasmids pHBT:STOP1-GFP and pHBT:GFP used in this study have been previously described⁵⁷. An effector or empty control plasmid (6 μg), and an internal control plasmid (pUBQ10:GUS; 2 μg) were co-transfected into 2 × 10⁵ protoplasts using the polyethylene glycol (PEG)-mediated transformation method. At least 4 independent biological replicates were set for each transformation, and then the protoplasts were harvested together for the isolation of chromatin. The sheared DNA was immunoprecipitated with anti-FLAG magnetic beads overnight at 4 °C. In addition, ChIP assays were also carried out using 10-day-old Col-0 and pBRM:BRM-FLAG/brm-3 plants (approximately 2 g) and Pierce agarose ChIP Kit (26156; Thermo Fisher Scientific), according to the manufacturer’s instructions. The sheared DNA was immunoprecipitated with anti-FLAG magnetic beads overnight at 4 °C. The enrichment of specific DNA fragments in the immune precipitant was determined by qPCR analysis of the recovered DNA using specific primers (Supplementary Data 2). The fold enrichment represents the immune-precipitation efficiency in the transgenic plants normalized to that in the Col-0 plants.

Formaldehyde-assisted isolation of regulatory elements (FAIRE)-qPCR analysis

To assess the effect of BRM disruption on the open chromatin in the genome of NRT1.1, FAIRE was performed using the whole seedlings of 10-day-old WT, brm-3, and brm-20 plants, according to the protocol described previously with modifications^60,61. Briefly, chromatins are cross-linked by 3% (final vol) formaldehyde treatment, followed by shearing chromatin by sonication, and reversing crosslinking. Then, a phenol/chloroform/isoamyl alcohol (PCI) extraction is performed, separating protein-free genomic DNA fragments into the aqueous phase and histone-DNA complexes to the interphase. The enrichment of specific DNA fragments in the recovered DNA was determined by qPCR analysis using specific primers (Supplementary Data 2). The relative enrichment of each DNA sample was first calculated using the delta cycle threshold (2^−∆∆Ct) method by calculating the ratio of FAIRE signal to input signal. Relative amplicon abundance represents the results in the mutants normalized to that in the Col-0 plants.

Transient expression assay

To performed transient expression assays in Nicotiana benthamiana leaves, three DNA segments, selected from NRT1.1 genomic region (P1 and P2) that containing the identified binding site of STOP1^5,14 and (P3) that containing the identified binding site of NLP7³² but not STOP or BRM, were individually inserted into the reporter vector pGreen0800-LUC using XhoI and BamHI restriction sites and using ClonExpress Ultra One Step Cloning Kit V3 (C117-01; Vazyme Biotech Co., Ltd, Nanjing, PRC). The primers used to generate the various clones are listed in Supplementary Data 2. In dual-luciferase reporter assays, reporter and effector constructs (35S:STOP1¹³ or 35S:NLP7) were co-transformed into young N. benthamiana leaves for 48 h. Firefly and Renilla luciferase activities were quantified using a dual-luciferase assay kit (Beyotime Biotechnology, China). Some infiltrated leaves were also sprayed with 1 mM luciferin (A600577; Sangon Biotech, Shanghai, China), after which the images of luminescence were captured using a cooled charge-coupled device (CCD) imaging apparatus (LB985; Berthold).

Measurement of NO₃ ⁻ uptake

For the assays of NO₃⁻ uptakes, 7-day-old seedlings were pre-treated in agar medium with a pH of 6.5 or 4.7, for 1 d. The plants were washed with 0.05 mM CaSO₄ for 1 min and then transferred to a medium in which KNO₃ was replaced with K¹⁵NO₃ (atom % ¹⁵N, 99%) for 10 min under the same conditions used for pre-culturing seedlings. The plants were washed with 0.05 mM CaSO₄ for 1 min, after which each plant was separated into shoots and roots, and the organs were dried at 65 °C for 3 d. The dried organs were weighed, and the roots and shoots of each plant were combined and ground into a fine powder. The samples were analyzed using an isotope ratio mass spectrometer (Isorime100; Elementar Analysensysteme, Hanau, Germany). The rate of ¹⁵NO₃⁻ uptake was calculated from the total ¹⁵N of the plant and the biomass of the roots.

Statistical analyses

Data were analyzed by one-way, two-way, or multi-way ANOVA with Tukey’s multiple comparison test.

Reporting summary

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

Data availability

All data are available in the main text or the supplementary materials. Source data are provided with this paper.

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Acknowledgements

We thank Dr. She Long Zhang (the Instrument Platform of College of Life Sciences, Zhejiang University) for his assistant in Confocal measurement, Prof. Ming Guang Lei (Hangzhou Normal University, Hangzhou, China), Dr. Thierry Desnos (CEA Cadarache), Prof. Yi Kun He (Capital Normal University, Beijing, China), Prof. Zhi Ye Wang (Zhejiang University), and Prof. Ai Wu Dong (Fudan University, Shanghai, China) for providing seeds. This study was supported by the National Key Research and Development Program of China (grant no. 2022YFA1303402 to Z.J.D. and S.J.Z.), the National Natural Science Foundation of China (grant no. 31970272 to Z.J.D., 32500240 to J.Y.Y., and 32201702 to W.H.T.), the Postdoctoral Fellowship Program of CPSF (grant no. GZC20232327 to J.Y.Y.), the China Postdoctoral Science Foundation (2023M743048 to J.Y.Y.), the Ministry of Education and Bureau of Foreign Experts of China (grant no. B14027) and the Fundamental Research Funds for the Central Universities, the Central Public-interest Scientific Institution Basal Research Fund (grant no, Y2023QC21 to W.H.T.).

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These authors contributed equally: Jia Yuan Ye, Wen Hao Tian.

Authors and Affiliations

State Key Laboratory of Plant Environmental Resilience, College of Life Science, Zhejiang University, Hangzhou, 310058, China
Jia Yuan Ye, De Rui Zhang, Qian Yi Yu, Ji Ming Xu, Chong Wei Jin, Shao Jian Zheng & Zhong Jie Ding
State Key Laboratory of Rice Biology and Breeding, China National Rice Research Institute, Hangzhou, 310006, China
Wen Hao Tian
Ningbo Key Laboratory of Agricultural Germplasm Resources Mining and Environmental Regulation, College of Science and Technology, Ningbo University, Ningbo, 315300, China
Wo Na Ding
College of Agronomy and Biotechnology, Zhejiang University, Hangzhou, China
Gui Xin Li
Agricultural Experimental Station, Zhejiang University, Hangzhou, 310058, China
Jing Ying Yan
Université Paris-Saclay, CNRS, INRAE, Univ Evry, Institute of Plant Sciences Paris-Saclay (IPS2), Orsay, 10 91405, France
Moussa Benhamed

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Jia Yuan Ye
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Contributions

S.J.Z., W.H.T., Z.J.D., and J.Y.Y. conceived and designed the experiments. J.Y.Y., W.H.T., and D.R.Z. performed the experiments. Q.Y.Y., J.M.X., G.X.L. and J.Y.Y. (Jing Ying Yan) assisted in plant material generation. J.Y.Y. and W.T.H. analyzed the data and generated the figures. W.N.D., C.W.J., and M.B. provided suggestions and revised the manuscript. J.Y.Y., Z.J.D., and S.J.Z. wrote the manuscript.

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Correspondence to Moussa Benhamed, Shao Jian Zheng or Zhong Jie Ding.

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Ye, J.Y., Tian, W.H., Zhang, D.R. et al. BRAHMA represses STOP1-NRT1.1 module to control plant rhizosphere alkalization and acid stress adaptation. Nat Commun 17, 3084 (2026). https://doi.org/10.1038/s41467-026-69905-z

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Received: 23 July 2025
Accepted: 13 February 2026
Published: 23 February 2026
Version of record: 31 March 2026
DOI: https://doi.org/10.1038/s41467-026-69905-z