FL7 is an ancient ABA-independent inhibitor of PP2C-As regulating plant stress responses

Li, Tianli; Yang, Zitong; Li, Guojun; Vries, Jan de; Li, Yuke; Li, Xianglan; Lu, Xu; Gu, Xinyi; Chen, Zhaojie; Zhao, Yang; Dou, Daolong; Shen, Danyu; Ai, Gan

doi:10.1038/s41467-025-66086-z

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Published: 15 December 2025

FL7 is an ancient ABA-independent inhibitor of PP2C-As regulating plant stress responses

Nature Communications volume 16, Article number: 11119 (2025) Cite this article

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Abstract

Clade A protein phosphatase 2Cs (PP2C-As) play crucial roles in plant stress responses. Although the ABA receptors PYLs inhibit PP2C-As in an ABA-dependent manner, other modulators of these phosphatases remain largely unknown. Here, we identify the FORKED-LIKE 7 (FL7) protein as a broad PP2C-A interactor that effectively suppresses PP2C activity through an ABA-independent, noncompetitive mechanism. By inhibiting PP2C-A activity, FL7 positively regulates osmotic tolerance and plant immunity in an ABA-independent manner. The N-terminal auxin canalisation (AC) domain of FL7 is required for its PP2C-As inhibitory activity. Further evolutionary analyses reveal that FL7 homologues containing an AC domain belong to an ancient family that emerged in a common ancestor of Klebsormidiophyceae algae and land plants. Genetic analyses indicate that algal FL7 homologues have a conserved function as PP2C-A inhibitors. Our study reveals an ABA-independent layer of PP2C-A modulation that regulates biotic and abiotic stress responses, which is likely conserved across a billion years of streptophyte evolution and predated the PYL-ABA regulation established in the common ancestor of land plants.

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Introduction

Owing to the climate crisis, plants face ever-increasing soil salinization and periods of drought, imposing osmotic stresses that adversely affect crop growth, yield, and resistance to disease^1,2. High soil salinity, in particular, heightens plant vulnerability to soil-borne pathogens. For instance, irrigation with saline water significantly compromises onion resistance to Fusarium oxysporum³, while tomato and chrysanthemum grown in high-salinity nutrient solutions exhibit increased disease severity caused by Phytophthora parasitica or P. cryptogea⁴. This phenomenon is partially attributed to the accumulation of the phytohormone abscisic acid (ABA), which antagonizes salicylic acid (SA), a vital defence hormone, under osmotic stress conditions^5,6. Consistently, ABA application induces disease susceptibility in different plants, such as Arabidopsis thaliana, tomato, and soybean, independent of the pathogen’s lifestyle^7,8. These findings highlight the crosstalk between plant osmotic stress and defence responses; however, the underlying mechanisms remain largely elusive, posing a considerable challenge to the development of climate-resilient crop varieties.

During evolution, intricate signalling mechanisms have emerged through which plants exhibit appropriate responses to withstand osmotic and pathogenic stress⁹. Central to these stress response pathways is the reversible phosphorylation of proteins, facilitated by the coordinated actions of protein kinases and protein phosphatases¹⁰. In particular, the clade A protein phosphatase 2 C (PP2C-A) family plays a pivotal role in regulating plant stress signalling networks¹¹ and might act as a hub linking plant osmotic and defence responses. Arabidopsis encodes 9 PP2C-As, including ABA INSENSITIVE 1/2 (ABI1/2), HYPERSENSITIVE TO ABA1/2 (HAB1/2), HYPERSENSITIVE GERMINATION 1/3 (AHG1/3), and HIGHLY ABA-INDUCED1/2/3 (HAI1/2/3). All of the enzymes are functional phosphatases and negative regulators of the plant osmotic stress response^12,13. For example, HAI1/2/3 single mutants exhibit increased proline and osmoregulatory solute accumulation at low water potential¹⁴, whereas AHG1/3 or ABI1/2 mutants exhibit increased drought tolerance¹⁵. PP2C-As also negatively regulate plant defence responses. HAI1 interacts with mitogen-activated protein kinases 3/6 (MPK3/6) and dephosphorylates them to suppress MPK3/6 activation, leading to weakened plant immunity¹⁶. ABI1 can also interact with and dephosphorylate MPK6¹⁷.

PP2C-As lack regulatory subunits^18,19; thus, their activities are precisely fine-tuned by those of other proteins. PYR/PYL/RCAR proteins (PYLs), which are important components of the ABA signalling pathway, are well known regulators of PP2C-As. In the absence of ABA, PP2C-As are associated with Snf1-related kinases 2 (SnRKs), including SnRK2.2/2.3/2.6, keeping them inactive by dephosphorylating their activation loops and blocking their catalytic clefts^20,21. PYLs are ABA receptors: in the presence of ABA, PP2C-As form complexes with PYL-ABA, and their activities are thus inhibited²². Such inhibition releases SnRK2s from suppression by PP2C-As, leading to the induced expression of many downstream effectors, such as RD29B²³. In the absence of ABA, several PYLs, such as PYL5, PYL6, PYL8, PYL9, and PYL10, can still interact with and inhibit PP2C-As, but the inhibition is weaker than that caused by ABA-bound PYLs²⁴.

In addition to PYLs, Enhancer of ABA Co-receptor 1 (EAR1) and the ROP GTPases ROP11 can enhance the activities of a subset of PP2C-As^25,26,27. Direct suppressors of PP2C-A phosphatase activity have also been identified. The RNA helicase RH8 and the haem-binding protein DOG1 physically associate with and suppress the activity of AHG3 and AHG1, respectively^28,29. Despite the numerous enzyme activity regulators identified for PP2C-As, these regulators merely function on several specific PP2C-As. Apart from PYLs, it remains unknown whether other broad-spectrum regulators target multiple PP2C-As.

The FORKED-LIKE (FL) protein family is characterised by an N-terminal auxin canalisation (AC) domain and a C-terminal pleckstrin homology-like (PH) domain^30,31. These proteins were initially discovered to be involved in leaf vein development by mediating asymmetric localisation of the auxin efflux protein PINFORMED1 (PIN1)³⁰. Recent studies have highlighted additional functions of FL proteins in plant biology. FL1 and FL3 are interaction partners of the blue light receptor cryptochrome (CRY2). CRY2 physically interacts with FL1 and FL3 to inhibit their ability to promote root elongation³². Our group found that FL7 transcripts are present across multiple plant tissues, including leaves, roots, stems, flowers, and siliques, with the protein localising in both nuclear and cytoplasmic compartments³³. Furthermore, we found that FL7 directly interacts with HAI1/2/3 and inhibits their phosphatase activity to prevent the dephosphorylation of MPK3/6, thereby promoting plant immunity³³. This discovery prompted us to investigate whether FL7, similar to PYLs, functions as a general enzyme inhibitor of PP2C-As.

In this study, we provide evidence that FL7 functions as a broad-spectrum inhibitor of PP2C-As in Arabidopsis and is thus a positive regulator of the plant ABA response and osmotic tolerance. Importantly, salt stress often impairs plant resistance to pathogens, but we found that FL7-overexpressing lines presented increased resistance to salt stress. In contrast to PYLs, FL7 inhibits PP2C-A activity in an ABA-independent and noncompetitive manner. Consistent with these findings, FL7 positively regulates the osmotic response independent of ABA. FL7 potently suppresses PP2C-A phosphatase activity by directly binding to its C-terminal phosphatase domain via its conserved N-terminal AC domain. Evolutionary analyses revealed that FL7 homologues emerged in Klebsormidiophyceae green algae and expanded across streptophyte lineages. The algal FL7 homologues, KnFL7 from Klebsormidium nitens and CbFL7 from Chara braunii, had PP2C-A inhibitory activity and exhibited functions that were similar to those of AtFL7. These results indicated that FL7 represents an ancient family of PP2C-A inhibitors that are conserved throughout green plants, predating PYLs, and might integrate plant biotic and abiotic stress responses.

Results

FL7 broadly interacts with PP2C-As and inhibits their phosphatase activity

We previously found that FL7 is an inhibitor of HAI1/2/3³³, prompting us to test the effect of FL7 on other PP2C-A members. First, we found that FL7 could interact with all of the other six PP2C-As in both yeast two-hybrid and coimmunoprecipitation assays, using HAIs as positive controls and a clade D PP2C (At5g02760) as a negative control (Fig. 1a,b). To confirm the natural interaction between FL7 and PP2C-As in planta, we immunoprecipitated the FL7 proteins from pFL7::gFL7-FLAG fl7 complementation lines (hereafter gFL7). ABI1 and ABI2 proteins could be detected in the precipitates using α-ABI1/ABI2 antibodies (Fig. 1c). Furthermore, we conducted pull-down assays to detect the direct interaction between FL7 and PP2C-As. We successfully purified the ABI1, ABI2, HAB2, and AHG3 proteins. Unlike the clade D PP2C control, all the purified PP2C-As displayed an affinity for FL7 in the pull-down assays (Fig. 1d). These results demonstrate that FL7 is associated with PP2C-As both in vivo and in vitro. Notably, PYL10 is a broad interactor of PP2C-As²⁴ and was also used as a positive control in the Y2H assay. We found that PYL10 interacts with all the PP2C-As except AHG1, whereas FL7 can interact with all of them (Fig. 1a).

**Fig. 1: FL7 is a general enzyme inhibitor of PP2C-As.**

We then tested whether FL7 can inhibit the activity of PP2C-As other than HAIs. The results revealed that FL7 inhibited between 23 and 56% of the activity of ABI1, ABI2, HAB2, and AHG3 but not that of the clade D enzyme (Fig. 1e), indicating that FL7 broadly inhibits PP2C-A activity. Given that the N-terminus of some PP2C-As, including ABI1 and AHG3, encompasses an autoinhibitory domain²⁵, we examined whether this region is required for FL7-mediated suppression. N-terminal truncated ABI1 (131–434) and AHG3 (106–398) mutants were generated, and their phosphatase activities were higher than those of the wild-type PP2C-As, indicating successful removal of the autoinhibitory domain (Supplementary Fig. 1a). FL7 still inhibited both the PP2C-A N-terminal truncation mutants (Supplementary Fig. 1a), demonstrating that FL7 directly suppresses PP2C-A activity independent of its autoinhibitory domain.

We then measured PP2C phosphatase activity among the wild-type, fl7 and OE-FL7 lines³³ in vivo (Fig. 1f). Our results demonstrated that compared with the wild-type plants, the FL7 overexpression lines presented significantly reduced PP2C activity, whereas the fl7 mutant presented the highest activity level (Fig. 1f). Collectively, these biochemical data confirm that FL7 acts as a broad-spectrum enzyme inhibitor of PP2C-As both in vivo and in vitro.

FL7 suppresses the dephosphorylation of MPK6 and SnRK2.6 mediated by PP2C-As

We further tested whether FL7 could suppress the PP2C-A-mediated dephosphorylation of its substrates. Phosphorylated MPK6 can be dephosphorylated by ABI1 (Supplementary Fig. 1b), which is consistent with a previous report¹⁷. In the presence of FL7, but not the GST control, MPK6 dephosphorylation by ABI1 was significantly restored (Fig. 1g). As a control, FL7 alone did not affect MPK6 phosphorylation (Supplementary Fig. 1c).

SnRK2s are well-known substrates of PP2C-As¹³. The phosphorylation level of SnRK2.6 greatly decreased in the presence of ABI1 (Fig. 1h, Supplementary Fig. 1d). However, this phenomenon was reversed by the addition of FL7 (Fig. 1h, Supplementary Fig. 1d), indicating that FL7 inhibited SnRK2.6 dephosphorylation by ABI1 in vitro. To test whether FL7 inhibits the PP2C-A-mediated dephosphorylation of SnRK2.6 in vivo, we conducted dual-luciferase reporter assays in snrk2.2/3/6 triple-mutant protoplasts (Supplementary Fig. 1e) because phosphorylated SnRK2.6 activates ABF2, which in turn induces RD29B expression²³. The results revealed that the expression of SnRK2.6 increased the pRD29B-LUC output, whereas the addition of ABI1 and AHG3 suppressed the chemiluminescence signal (Supplementary Fig. 1f). PYL10 was used as a positive control in this assay, and the addition of PYL10 restored the signals inhibited by ABI1 and AHG3 (Supplementary Fig. 1f).

FL7 positively regulates the plant osmotic stress response

To investigate the function of FL7 as an inhibitor of PP2C-As, we tested the ABA sensitivity of FL7-related genotypes on the basis of the antagonistic effect between total PP2C-A activity and the ABA response²⁵. In the absence of ABA, compared with the wild-type plants, the fl7 mutants and FL7 overexpression lines (hereafter OE-FL7) presented no obvious differences in terms of the germination rate or primary root length (Fig. 2a, Supplementary Fig. 2a). However, compared with the wild-type plants, the ABA treatment increased the fl7 mutant germination rate and primary root growth, whereas the OE-FL7 seedlings displayed ABA hypersensitivity (Fig. 2a, Supplementary Fig. 2a). Similarly, ABA-induced stomatal closure was impaired in the fl7 mutants but enhanced in the OE-FL7 lines (Fig. 2b). These results indicated that FL7 positively regulates plant ABA responses, probably through inhibition of total PP2C-A activity.

**Fig. 2: FL7 positively regulates plant osmotic stress response.**

PP2C-As are vital negative regulators of the plant osmotic response³⁴. We thus explored the role of FL7 in plant osmotic tolerance. Under normal conditions, all the Arabidopsis lines exhibited similar growth. After 20 days of drought, the fl7 mutant displayed the lowest survival rate, whereas the OE-FL7 lines showed significantly increased survival (Fig. 2c), demonstrating that FL7 positively regulates drought tolerance. Similarly, water loss from detached leaves was also more rapid in the fl7 mutant and weakened in the OE-FL7 lines (Supplementary Fig. 2b). To obtain a genome-wide view of gene expression changes in the fl7 mutant and OE-FL7 lines in response to osmotic stress, we conducted RNA sequencing (RNA-seq) analysis on Col-0, fl7 and OE-FL7 lines treated with 300 mM mannitol. After 12 h of treatment, 2035, 2823 and 3314 differentially expressed genes (DEGs) ( | Log₂(fold change)| >= 1, P < 0.05) were detected in Col-0, fl7 and OE-FL7 seedlings, respectively (Supplementary Data 1). The overlapping pattern of DEGs between Col-0, fl7, and OE-FL7 seedlings suggested a common osmotic response mechanism among these lines (Fig. 2d). The k-means clustering of all DEGs between Col-0, fl7, and OE-FL7 seedlings resulted in six clusters with distinct expression patterns (Supplementary Fig. 3). Cluster 3 included genes whose expression was strongly induced by mannitol in the OE-FL7 line but weakly induced in both the Col-0 and fl7 lines. The genes in Cluster 6 exhibited the strongest mannitol-induced expression in the OE-FL7 line, moderate induction in Col-0, and the weakest induction in the fl7 mutant (Supplementary Fig. 3). Gene Ontology (GO) enrichment analysis of the DEGs in Clusters 3 and 6 revealed that these genes were highly enriched in potential abiotic stress response-related GO terms such as “response to abiotic stimulus” and “response to stress” (Supplementary Fig. 3). Moreover, these clusters are enriched in many plant osmotic response marker genes, such as ABI1/2, RD29B and RAB18³⁵ (Supplementary Fig. 3). These marker genes generally exhibited the strongest mannitol-induced expression in the OE-FL7 line, moderate induction in Col-0, and the weakest induction in the fl7 mutant (Fig. 2e). These results indicated that FL7 positively regulates the plant osmotic response.

Enhanced pathogenic resistance of FL7-overexpressing lines is not impaired by salt stress

We previously reported that FL7 is a positive regulator of plant immunity³³. Combined with the data presented here, these findings reveal that FL7 is a positive hub regulator of both the osmotic response and the defence response, indicating that FL7 might be a promising genetic resource for the development of resistant plants suitable for saline soils. We first tested the role of FL7 in plant salt tolerance and found that FL7 promotes plant tolerance to salinity stress caused by NaCl through assays of electrolyte leakage (Fig. 2f). Activation of SnRK2s is a central step in the plant response to salt stress, and we found that NaCl-induced SnRK2.2, SnRK2.3, and SnRK2.6 phosphorylation was reduced in the fl7 mutant but increased in the OE-FL7 lines (Supplementary Fig. 4a). To test whether FL7 responded to salt stress, we analysed its protein accumulation level in OE-FL7 and gFL7 plants after NaCl treatment. The FL7 protein level increased with treatment (Supplementary Fig. 4b). These results indicated that FL7 responses to and positively regulates plant salt tolerance.

We next tested whether FL7 could promote plant immunity under salt stress. Four-week-old Arabidopsis plants were irrigated with water that contained 100 mM NaCl to simulate soil salinization, and the leaves were detached for the inoculation assay at 24 h post-treatment (Supplementary Fig. 4c). In the absence of NaCl treatment, the fl7 mutant was more susceptible, whereas the OE-FL7 line was more resistant to Phytophthora capsici (Fig. 2g), supporting FL7 as a positive plant immune regulator. After NaCl treatment, the susceptibility of the wild-type, fl7 mutant and gFL7 complementation lines to P. capsici significantly increased (Fig. 2g), which is consistent with previous findings that soil salinization increased disease severity caused by Phytophthora parasitica and P. cryptogea⁴. Interestingly, no obvious lesion area changes could be detected in the OE-FL7 lines after mock or NaCl treatment, indicating that the degree of immune weakness caused by NaCl treatment greatly decreased in the OE-FL7 plants (Fig. 2g).

Genetic analysis of fl7 with the pp2c-As mutants

To further analyse the genetic relationship between FL7 and PP2C-As, we knocked out FL7 in the abi1 hab1 pp2ca triple mutant using CRISPR-Cas9, generating an abi1 hab1 pp2ca fl7 quadruple mutant (Supplementary Fig. 4d). The abi1 hab1 pp2ca triple mutant is sensitive to ABA during seed germination and root growth (Fig. 3a, b), which is consistent with previous reports¹⁵. Although fl7 showed reduced ABA sensitivity compared with wild-type, the abi1 hab1 pp2ca fl7 quadruple mutant exhibited a phenotype comparable to that of the abi1 hab1 pp2ca triple mutant (Fig. 3a, b). Similarly, a drought tolerance analysis revealed that the abi1 hab1 pp2ca fl7 quadruple mutant displayed a phenotype comparable to that of the abi1 hab1 pp2ca triple mutant, with both showing enhanced drought tolerance (Fig. 3c). These results indicate that FL7 functioning is dependent mainly on PP2C-As.

FL7 inhibits PP2C-As in an ABA-independent and noncompetitive manner

ABA promotes the inhibition of PP2C-As by PYLs²². However, our phosphatase activity indicated that the inhibitory function of FL7 is independent of ABA (Fig. 1e)—in contrast to that of canonical PYLs. To further test whether ABA is required for FL7 function, a microscale thermophoresis binding assay was conducted by using PYL10 as a positive control. The results revealed an interaction between ABA and PYL10 but no binding affinity for ABA for the FL7 and GST control (Fig. 4a). We then tested whether ABA influences the interaction between FL7 and PP2C-As. Consistent with previous findings²², ABA stimulated PYL10-ABI1 complex formation (Fig. 4b). In contrast, ABA did not affect FL7-ABI1 associations (Fig. 4b). We next investigated whether ABA regulates FL7 inhibition of PP2C-A activity. Enzymatic analyses demonstrated that ABA bolstered PYL10-mediated but not FL7-mediated inhibition of ABI1 (Fig. 4c). Similar ABA-independent inhibition activity was observed for FL7 when ABI2 and AHG3 were used (Supplementary Fig. 5a, b). Furthermore, ABA did not enhance the ability of FL7 to suppress ABI1-mediated dephosphorylation of SnRK2.6 (Fig. 1h, Supplementary Fig. 1d). These results indicated that the inhibitory activity of FL7 on PP2C-As is independent of ABA.

**Fig. 4: FL7 inhibits PP2C-As through an ABA-independent noncompetitive mechanism.**

To elucidate the biochemical mechanisms underlying the inhibitory activities of FL7 and PYL10, we performed enzyme kinetic analyses (Fig. 4d). Competitive inhibitors compete with substrate(s) for the active site of the enzyme, and the inhibition is reversible upon substrate saturation. In contrast, noncompetitive inhibitors alter active site conformation or chemistry to reduce substrate binding/catalysis independently of substrate levels (Supplementary Fig. 5c)³⁶. Consistently, ABA did not increase FL7 inhibition in this assay (Fig. 4d). In contrast to the reversible inhibition of ABI1 by PYL10 upon substrate saturation despite the presence of ABA, FL7-mediated ABI1 suppression was unresponsive to both increased substrate availability and increased ABA treatment (Fig. 4d). Identical patterns also occurred for AHG3 (Supplementary Fig. 5d), corroborating a noncompetitive inhibition mechanism for FL7.

FL7 enhances plant osmotic stress tolerance through an ABA-independent pathway

To investigate whether FL7 regulates the plant osmotic response independently of ABA, we crossed the ABA-deficient aba1-1 mutant with fl7 to generate the fl7 aba1-1 double mutant (Supplementary Fig. 5e, f). We found that, in the absence of ABA, the stomata of aba1-1 were more open than those of fl7 and Col-0 were, while the stomatal aperture of the fl7 aba1-1 double mutant was the largest (Fig. 4e). Drought tolerance assay revealed that compared with the aba1-1 single mutant, the fl7 aba1-1 double mutant had lower survival rates (Fig. 4f, Supplementary Fig. 5g). These results further support that FL7 suppresses PP2C-A activity and positively regulates the plant osmotic response in an ABA-independent manner.

PYLs function as ABA receptors and broad suppressors of PP2C-As²². To examine the relationship between FL7 and PYLs, we employed ABA ANTAGONIST1 (AA1), a broad-spectrum antagonist that blocks all PYR/PYL-PP2C interactions³⁷ (Fig. 4g). We analysed the germination rates of Col-0 and fl7 mutant seeds grown on 1/2 MS plates supplemented with either ABA alone or ABA plus AA1. The addition of AA1 significantly increased the germination rate of Col-0, indicating successful inhibition of PYL activity (Fig. 4h). Notably, the germination rate of fl7 + AA1 was higher than that of Col-0 + AA1 (Fig. 4h), suggesting that FL7 continues to regulate the PP2C pathway even when PYL activity is blocked. In addition, we found that the presence of both PYL and FL7 led to a stronger inhibition of ABI1 phosphatase activity compared with either component alone, indicating an additive effect (Supplementary Fig. 5h). Collectively, these data support that FL7 and PYLs function in parallel pathways.

FL7 inhibits PP2C activity through an N-terminal auxin canalisation domain

The FL7 protein contained an N-terminal auxin canalisation domain (AC) and a C-terminal pleckstrin homology-like domain (PH) (Fig. 5a). To investigate which domain of FL7 is critical for the inhibitory activity on PP2C-As, we generated FL7 truncation mutants (FL7N and FL7C) (Fig. 5a). ABI1 and AHG3 were selected as representatives of PP2C-As to test their interactions with FL7 truncation mutants. The results showed that the AC domain, but not the PH domain, sufficiently interacted with the phosphatase domain of the tested PP2C-As in a Y2H assay (ABI1N for the autoinhibition domain; ABI1C for the phosphatase domain) (Fig. 5a, Supplementary Fig. 6a). Similarly, the AC domain was sufficient to suppress the activities of ABI1 and AHG3 (Fig. 5b, Supplementary Fig. 6b), and this inhibitory activity was not enhanced by ABA (Fig. 5c, Supplementary Fig. 6c). Furthermore, we added the AC domain or the PH domain of FL7 under the control of the native promoter of FL7 to the fl7 mutant line (Supplementary Fig. 6d). The AC domain, but not the PH domain, completely restored the fl7 mutant phenotype, while overexpression of the AC domain mimicked the phenotype of the OE-FL7 lines (Fig. 5d–f, Supplementary Fig. 6e–g). Compared with the OE-FL7 lines, the protein extracts of OE-FL7N lines also exhibited impaired PP2C activity (Fig. 1f). These results indicated that the AC domain is sufficient for the suppression of PP2C activity.

**Fig. 5: FL7 inhibits PP2Cs activity through an N terminal auxin canalisation domain.**

FL7 emerged in an ancient streptophyte

When did FL7 evolve? To investigate the conservation of FL7, we performed comparative genomic and phylogenetic analyses of the AC domain in the genomes of 17 representative plant species that cover a broad diversity of plants (Fig. 6a; Supplementary Data 2). Using a Markov model of the AC domain generated by the Pfam database, we identified a total of 32 FL7 homologues with the AC domain in the genomes of 9 plants (Fig. 6a; Supplementary Data 3 and 4). The number of FL7 homologues we identified in Arabidopsis aligns with previous reports³⁰, bolstering the identification stability of our approach. In addition to the land plant clade, homologues of FL7 were found in the streptophyte algal families Klebsormidiophyceae and Charophyceae but not in the phragmoplastophytic Coleochaetophyceae and Zygnematophyceae or the clade of Mesostigmatophyceae and Chlorokybophyceae (Fig. 6a). We performed a large-scale phylogenetic analysis of these proteins with the AC domain. We found that plants formed a monophyletic group with high support values (Fig. 6a). The copy number of FL7 homologues is low in algal and bryophyte species but is expanded in vascular plants (Fig. 6a, b), indicating that FL7 homologues are monophyletic.

**Fig. 6: FL7 is conserved across streptophyte.**

PYLs originated in the common ancestor of Zygnematophyceae and land plants³⁸ and were likely gained by horizontal gene transfer from soil bacteria³⁹. Interestingly, FL7 homologues were identified in Klebsormidiophyceae, suggesting that they emerged in a last common ancestor (LCA) of Klebsormidiophyceae and Phragmoplastophyta, which likely lived more than a billion years ago⁴⁰. These findings indicate that FL7 homologues might be more ancient PP2C inhibitors than PYLs are. To further study FL7 homologues in streptophyte algae, we performed similarity searches and phylogenetic analyses within six charophyte transcriptomes^41,42. We found that FL7 homologues are present in representatives of two other charophyte genera: Nitella, another Charophyceae, and, importantly, Coleochaete, a Coleochaetophyceae (Supplementary Data 5). These findings further underscore the emergence of FL7 homologues in the LCAs of Klebsormidiophyceae and Phragmoplastophyta. Since the absence of one gene in the transcriptome of one species does not guarantee its absence in that species⁴³, it might be that FL7 homologues are even more widespread in streptophytes. Our results further demonstrate that the AC domain is widely present in streptophyte algae. Although some plant FL7 homologues lack a PH domain (Fig. 6a; Supplementary Data 4), these proteins cluster with those with PH domains in the phylogenetic tree (Fig. 6a), indicating that they might have arisen from AC-PH proteins by deleting PH. Taken together, our comparative genomic analyses provide evidence for the deep evolutionary origin of FL7 homologues in a streptophyte predating PYLs (Fig. 6b).

FL7 function is conserved across a billion years of streptophyte evolution

PYL was an evolutionary innovation of a common ancestor of Zygnematophyceae and land plants^39,44. Interestingly, our evolutionary analysis revealed that compared with PYLs, FL7 homologues are more ancient PP2C inhibitors. To investigate their conserved function across streptophytes, we synthesised FL7s from the two streptophyte algae Klebsormidium nitens and Chara braunii. Y2H and pull-down assays revealed that, like AtFL7, algal FL7s also interact with AtABI1 (Fig. 7a, b). Furthermore, algal FL7 inhibited the activity of AtABI1 (Fig. 7c), indicating that the function of algal FL7 is conserved with AtFL7. We constructed transgenic plants in which the algal FL7s were under the control of the AtFL7 promoter and introduced them into the fl7 mutant (Supplementary Fig. 7a, b). Compared with the wild-type plants, the complemental lines presented a similar osmotic response (Fig. 7d, e). Thus, our results indicated that FL7 is a functionally conserved PP2C-A inhibitor across a billion years of streptophyte evolution.

**Fig. 7: FLs are functionally conserved in Klebsormidiophyceae and Charophyceae.**

Discussion

To survive in unfavourable environments, plants activate diverse adaptive processes to prevent osmotic stress and pathogen colonization, among which PP2C-As are key modulators⁴⁵. Here, we report that a broad-spectrum PP2C-A interactor, FL7, functions as a direct inhibitor of PP2C-A activity through its N-terminal AC domain. Interestingly, the function of FL7 inhibition differs from that of PYLs in two ways: FL7 inhibition of PP2C-A is independent of ABA and occurs in a noncompetitive manner. Furthermore, FL7 positively regulates osmotic tolerance in an ABA-independent manner. The FL7-overexpressing lines were highly resistant to pathogens that could not be impaired by osmotic stress (Fig. 7f). Finally, we found that FL7 is a conserved PP2C inhibitor that emerged more than a billion years ago (Fig. 6b). The functions of FL7 homologues from streptophyte algae are similar to those in Arabidopsis.

Soil salinity is an increasingly severe global problem as a result of improper irrigation practices and climate change. The effects of such environmental variables on plants can have negative impacts on plant disease development¹. Identifying key regulators that integrate osmotic stress and defence responses could aid in the development of resistant crops adapted to climate change. Here, we demonstrate that overexpression of FL7, a PP2C-As inhibitor, facilitates plant adaptation to both abiotic and biotic stress. More importantly, FL7 overexpression prevents the impairment of plant immunity by salt stress (Fig. 2g), indicating that FL7 might be a good genetic resource for improving the plant defence response under saline conditions. However, as endogenous FL7 also accumulated under salinity but its resistance was impaired by salinity, we assumed that the high level of FL7 accumulation prior to salinity stress might be critical for conferring resistance to pathogens under salinity. Future research should focus on fine-tuning FL7 protein levels to promote pathogen resistance under salinity but avoid potential side effects. Such findings highlight the strategy of inhibiting negative hub regulators, such as PP2C-As, in plant responses to biotic and abiotic stresses, which might aid in the development of crops adapted to fluctuating environments caused by climate change.

Interestingly, FL7 inhibited PP2C-As in an ABA-independent manner. In addition to ABA-dependent pathways, ABA-independent pathways are important adaptive mechanisms in plants^13,46. For example, the ahg1-1 mutant has no ABA-related phenotype in adult plants⁴⁷. Such a function of AHG1 might result from its specific interaction surface with PYLs, leading it to interact only with RCAR1-3¹². Although ABA-bound RCAR1-3 interacts with AHG1, it only weakly inhibits AHG1 activity¹². Unlike PYLs, DOG1 can interact with AHG1 and suppress its activity in parallel with the ABA pathway⁴⁸. Here, we found that FL7 inhibits PP2C-As in an ABA-independent manner and that FL7 also interacts with AHG1 (Fig. 1a), indicating that FL7 interacts with PP2C-As in a different way than PYLs do. This assumption could be further proven by the observation that FL7 inhibits PP2C-As and osmotic tolerance in a noncompetitive manner, which is also different from the activity of PYLs (Fig. 4d). Importantly, mutation of FL7 in the ABA-deficient aba1-1 mutant further weakened the impaired osmotic tolerance of aba1-1 (Fig. 4f), indicating that FL7 is involved in the ABA-independent osmotic response.

How plants adapted to living on land is a long-standing question. The evolution of the PP2C-A-regulated pathway is considered a key step in plants’ ability to adapt to terrestrial environments⁴⁹. PP2C-As show high functional conservation among all streptophyte species, including algae such as the Klebsormidiophyceae and Charophyceae^50,51. However, PYLs originated in the common ancestor of Zygnematophyceae and embryophytes and were obtained by horizontal gene transfer from soil bacteria^39,44. This raises the question of how ancient algae such as K. nitens and C. braunii, which lack a PYL system, regulate PP2C-A activity, causing them to suffer from osmotic stress. Here, we showed that FL7 is a conserved PP2C-A inhibitor throughout Streptophyta and that FL7 homologues from K. nitens and C. braunii also exhibit PP2C-A inhibition ability (Fig. 7a–e), indicating that FL7 homologues are ancient PP2C-A regulators that might have been used by early algae to regulate PP2C-A activity and adapt to osmotic stress. This assumption could be proven by investigating the biological function of FL7 homologues in these algae in the future. Furthermore, the FL7 homologues expanded during plant terrestrialization (Fig. 6a, b). Notably, treatment of the fl7 mutant with the general PYL inhibitor AA1 further impaired ABA sensitivity, suggesting that FL7 continues to regulate the PP2C pathway even when PYL activity is blocked (Fig. 4h). These findings indicate that the FL7-PP2C module may represent another important regulatory branch parallel to the ABA-PYL pathway in plant terrestrial adaptation and might help plants regulate ABA-independent PP2C-A regulatory pathways.

The FL family expanded during land plant evolution. In Arabidopsis, FL7 is one of nine FKD1/FL proteins. The other FLs also contain an AC domain (Fig. 6a) and thus might also be PP2C-A inhibitors. There might be functional redundancy between FLs. FLs may interact with PP2C-As to form a complex network, thus potentially facilitating responses to a plethora of physiological and developmental processes and environmental stresses. Unravelling this network will require considerable future investigation.

Collectively, our data indicate that FL7 homologues represent an ancient PP2C-A inhibitor family conserved throughout green plants and integrate plant biotic and osmotic responses.

Methods

Plant materials and growth conditions

Arabidopsis plants were grown and maintained in plant growth chambers at an ambient temperature of 23 °C under a 16-h light/8-h dark photoperiod. Light was provided by white fluorescent bulbs with an intensity of ~120 μmol/m²/s. All Arabidopsis lines used in this study were from a Col-0 accession. T-DNA insertion mutants were obtained from Arashare (https://www.arashare.cn/index/).

We previously obtained fl7, OE-FL7, gFL7, abi1 hab1 pp2ca and snrk2.2/3/6 triple mutants^33,52,53. The 1000 bp FL7 promoter (pFL7) was amplified from Col-0 genomic DNA. The sequences of KnFL7 and CbFL7 were synthesised by a biological company. The coding sequences of FL7 and PYL10 were subsequently cloned via RT‒PCR. To generate complementation transgenic lines expressing FLAG-tagged pFL7:FL7N, pFL7:FL7C, pFL7:KnFL7 and pFL7:CbFL7, DNA fragments encoding these sequences were fused to a C-terminal 3 × FLAG tag and cloned and inserted into the pCambia1300 vector. To generate transgenic lines overexpressing FLAG-tagged FL7N, the sequence encoding FL7N was fused to a C-terminal 3 × FLAG tag and cloned and inserted into the pCambia1300 vector. All the constructs were subsequently introduced into Agrobacterium (Agrobacterium tumefaciens) strain GV3101 for plant transformation using the floral dip method. Positive transformants were recovered by antibiotic selection and confirmed by genomic PCR. The abi1 hab1 pp2ca fl7 quadruple mutant was generated by knocking out FL7 in the abi1 hab1 pp2ca triple mutant using CRISPR-Cas9. To generate the fl7 aba1-1 double mutant, an fl7 T-DNA insertion mutant (SALK_07717C) was crossed with an aba1-1 T-DNA insertion mutant (SALK_027326C). The primers used in this study are listed in Supplementary Data 6.

For the root growth assay, the corresponding seeds were germinated on 1/2 MS medium. After incubation at 4 °C for 3 days, the seeds were transferred to a greenhouse with a 16-h light/8-h dark cycle for an additional 3 days. Seedlings with similar root lengths were then transferred to 1/2 MS medium supplemented with various concentrations of ABA and vertically grown for an additional 5 days. The root lengths were measured using ImageJ software.

For the germination assays, seeds were sown on 1/2 MS medium supplemented with ABA and incubated at 4 °C for 3 days. Unless otherwise stated, a concentration of 0.2 µM ABA was used for all germination assays. The plates were then transferred to a greenhouse and grown for an additional 7 days before determining the seed germination ratio. For the experiment shown in Fig. 4h, a concentration of 0.3 µM ABA was employed, as this assay was conducted following the relocation of our laboratory, during which it was determined that 0.2 µM ABA was insufficient to effectively suppress seed germination.

Drought-stress tolerance

To assess drought tolerance, the seedlings were transferred to plastic pots, grown for 3 weeks under short-day conditions, and then subjected to drought conditions by withholding water from the plants for 20 days. The drought-induced phenotypes were photographed with a camera. The survival rates of each genotype were analysed at 2 days after rewatering.

Stomatal aperture measurements

For the stomatal aperture assay, leaves were collected from 5-week-old plants and incubated in MES buffer (10 mM MES-KOH (pH 6.15), 10 mM KCl, and 50 µM CaCl₂) under light for 3 h at 23 °C to open the stomata. To examine ABA-induced stomatal closure, 20 µM ABA was added to the MES buffer, which was subsequently incubated for an additional 2 h under the same conditions. In each individual sample, at least 80 stomata were randomly observed, and images were obtained with a microscope. The stomatal aperture was measured using ImageJ software.

Measurement of water loss

Three-week-old seedlings grown under short-day conditions (16-h light/8-h dark) were used for water loss measurements. The whole rosettes were cut from the plants and weighed at the indicated points.

Ion leakage assay

Ion leakage induced by cell death was analysed as previously described⁵⁴. In brief, three 9-mm leaf discs were rinsed three times with distilled water and then soaked in 9 mL distilled water for 3 h at room temperature. The conductivity of the solution was measured using a conductivity meter to generate S1. The leaf discs were then boiled for 20 min, after which the conductivity was measured again to determine the value of S2. Electrolyte leakage was calculated as the percentage (%) of “S1”/“S2”. The assay was repeated three times.

Immunoblot assay

To extract proteins from plant materials, leaves were frozen in liquid nitrogen and ground to a fine powder. Extraction buffer (50 mM HEPES, 150 mM KCl, 1 mM EDTA, and 0.1% [v/v] Triton X-100; pH 7.5) containing 1 mM DTT and protease inhibitor cocktail (Sigma) was used for protein extraction. Anti-GFP antibody (1:5000; #M20004; Abmart) and HRP-conjugated anti-mouse IgG (1:10,000, Sigma‒Aldrich) secondary antibody were used for immunoblotting protein with a GFP tag. mAb-HRP-DirecT anti-FLAG antibody (1:5000; #M185-7; MBL) was used for immunoblotting proteins with a FLAG tag. mAb-HRP-DirecT anti-HA antibody (1:5000; #M180-7; MBL) was used for immunoblotting proteins with an HA tag.

Yeast two-hybrid assay

To detect the interaction between FL7 and PP2C-As, the cDNA fragments of FL7 and PP2C-As were amplified by PCR and cloned and inserted into the pGBKT7 and pGADT7 vectors, respectively. The bait and prey constructs were cotransformed into the yeast strain AH109. Successfully transformed colonies were selected on yeast SD medium lacking Trp and Leu. Transformants were subsequently transferred to selective SD medium lacking Trp, Leu, and His for growth analysis. Photographs were taken after 4 days of incubation at 30 °C.

Co-IP assay

Arabidopsis protoplasts were transfected with the indicated plasmid combinations and control constructs, and transfection was performed as described above. Immunoprecipitation with anti-FLAG antibodies was carried out as previously described⁵⁵. Total protein and immunoprecipitates were separated by SDS–PAGE and detected by immunoblotting.

Protein purification

Proteins with different tags were expressed in Rosetta (DE3) Escherichia coli cells growing in LB medium supplemented with 0.5 mM IPTG for 18 h at 18 °C. The culture was then collected by centrifugation at 6000 × g for 10 min at 4 °C. For His-tagged proteins, the pellet was resuspended in lysis buffer (50 mM NaH₂PO₄, 30 mM NaCl, 10 mM imidazole (pH 8.0), 1 mM PMSF, and protease inhibitor cocktail [Abmart, A10004]) and then sonicated for 10 min. After centrifugation for 30 min at 7000 g and 4 °C, the supernatant was incubated with Ni Sepharose (GE) resin for 3 h at 4 °C. The resin was subsequently washed three times with washing buffer (50 mM NaH₂PO₄, 30 mM NaCl, and 50 mM imidazole; pH 8.0) to remove nonspecifically bound proteins. The His-tagged proteins were eluted from the resin with elution buffer (50 mM NaH₂PO₄, 30 mM NaCl, and 250 mM imidazole, pH 7.4). For the GST-tagged proteins, the pellet was resuspended in phosphate-buffered saline (PBS) (150 mM NaCl, 10 mM Na₂HPO₄, 2 mM KH₂PO₄, 2.7 mM KCl (pH 7.4), 1 mM PMSF, and inhibitor cocktail) and then sonicated for 10 min. After centrifugation for 30 min at 7000 g and 4 °C, the supernatant was incubated with glutathione Sepharose 4B (GE) resin for 3 h at 4 °C. The resin was subsequently washed three times with PBS to remove nonspecifically bound proteins. The GST-tagged proteins were eluted from the resin with GSH buffer (50 mM Tris-HCl, pH 8.0, and 10 mM GSH). For MBP-tagged proteins, the pellet was resuspended in lysis buffer (20 mM Tris–HCl (pH 7.4), 200 mM NaCl, 1 mM EDTA, 1 mM PMSF, and inhibitor cocktail) and then sonicated for 10 min. After centrifugation for 30 min at 7000 g and 4 °C, the supernatant was incubated with amylose resin (NEB) for 3 h at 4 °C. The resin was subsequently washed three times with lysis buffer to remove nonspecifically bound proteins. The MBP-tagged proteins were eluted from the resin with elution buffer (20 mM Tris-HCl (pH 7.4), 200 mM NaCl, 1 mM EDTA, and 10 mM maltose). To measure their concentrations, the purified proteins were mixed with Quick Start Bradford Dye Reagent (Bio-Rad), and the absorbance was measured at 595 nm. Protein concentrations were calculated on the basis of a standard curve with bovine serum albumin (BSA) as a substrate.

Protein pull-down assay

The coding sequences of the genes of interest were subsequently cloned and inserted into the MBP vector or the pGEX-6P vector. Recombinant MBP-PP2C, GST, and GST-FL7 proteins were purified as described above. Purified GST or GST-FL7 (10 mg) in 2 mL of lysis buffer (150 mM NaCl, 10 mM Na₂HPO₄, 2 mM KH₂PO₄, 2.7 mM KCl (pH 7.4), 1 mM PMSF, and inhibitor cocktail) was incubated with glutathione beads for 2 h at 4 °C. After a brief centrifugation at 1000 × g for 3 min at 4 °C, the buffer was removed, and 2 mg of recombinant MBP-PP2C protein was added to the resin in 2 mL of lysis buffer. The tube was rotated at 4 °C for 2 h for protein binding. The resin was then washed five times with PBS to remove nonspecifically bound protein, resuspended in 50 mL of PBS and 10 mL of SDS loading buffer, and boiled for 5 min. After centrifugation at 12,000 × g for 1 min at 22 °C, the supernatant was subjected to immunoblotting analysis. Immunoblot assays were conducted with anti-GST (1:5000; #M20007; Abmart) or anti-MBP (1:5000; #M20001; Abmart) antibodies and HRP-conjugated anti-mouse IgG (1:10,000; Sigma‒Aldrich) as the secondary antibody.

MST assay

The interactions between ABA and recombinant GST-FL7, GST-PYL10, and GST were measured by MST (microscale thermophoresis). MST assays were performed as previously described⁵⁶.

First, the purified GST, GST-FL7, and GST-PYL10 sample proteins were exchanged for buffer with labelling buffer (130 mM NaHCO₃, 50 mM NaCl, pH 8.2–8.3) and then labelled with dye in assay buffer using the Protein Labelling Kit RED-NHS 2nd Generation (Cat# MO-L011). ABA at a range of concentrations (0.3 µM to 10 mM) was incubated with 2.5 µM labelled protein for 30 min at room temperature, after which the samples were loaded onto NanoTemper glass capillaries (NanoTemper Technologies), and MST was performed. The experimental data were processed, and the Kd values for ABA-FL7, ABA-PYL10, and ABA-GST binding were calculated through MO. Affinity Analysis software.

In vitro dephosphorylation of MAPK

The Escherichia coli strain Rosetta (DE3) was used for the production of recombinant proteins. To prepare phosphorylated MPK6, 5 µg of recombinant purified MBP-tagged kinase-inactive MAPK was incubated with 1 µg of GSTMKK4DD at 30 °C for 1 h in kinase buffer (25 mM Tris–HCl (pH 7.5), 1 mM DTT, 10 mM MgCl₂, and 200 µM ATP). The kinase reaction was terminated by removing ATP from the reaction using an Amicon Ultra30K centrifugation unit (Millipore). To test the phosphatase activity of ABI1, phosphorylated MAPK (500 ng) was incubated with MBP or MBP-ABI1 (2 µg) at 30 °C for 15 min in phosphatase buffer (25 mM Tris-HCl (pH 7.5), 1 mM DTT, and 10 mM MgCl₂). MAPK was detected by immunoblotting as described above. To test whether FL7 inhibits the phosphatase activity of ABI1, phosphorylated MAPK (500 ng) and MBP-ABI1 (2 µg) were incubated with GST or GST-FL7 at 30 °C for 15 min in phosphatase buffer (25 mM Tris-HCl (pH 7.5), 1 mM DTT, and 10 mM MgCl₂).

Measurement of PP2C-A activity

In vitro PP2C-A activity was measured using a Ser/Thr Phosphatase Assay Kit (Promega, V2460). In brief, the recombinant proteins were purified with buffers that did not contain phosphate. To measure phosphatase activity, different combinations of recombinant purified FL7 and PP2Cs were mixed with 5 mL of 1 mM phosphopeptide and 10 mL of PP2C reaction buffer (250 mM imidazole (pH 7.2), 1 mM EGTA, 25 mM MgCl₂, 0.1% [v/v] β-mercaptoethanol, and 0.5 mg/mL BSA) in the wells of a microtiter dish and then incubated at 25 °C for 15 min. The reactions were stopped by the addition of 50 mL of molybdate dye/additive mixture, after which the plate was incubated at room temperature for 15 min. The absorbance was measured at 630 nm in a microplate reader (Molecular Devices, SpectraMax M5). Michaelis–Menten plots in GraphPad Prism 9.0 were used to calculate the kinetic parameters.

To measure PP2C activity in vivo, total proteins (0.1 g) from the wild type or fl7, OE-FL7, or OE-FL7N mutants were extracted using phosphatase storage buffer (20 mM Tris–HCl (pH 7.5), 20 mM KCl, 1 mM EDTA, 1 mM EGTA, 10 mM DTT, 0.5% [v/v] Triton X-100, and 50% [v/v] glycerol) with a protease inhibitor cocktail (Abmart, A10004), after which the homogenised lysate was centrifuged at 100,000 × g for 1 h at 4 °C. The supernatant was filtered through Sephadex G-25 resin to remove endogenous phosphates, after which the lysate was used to measure PP2C activity as described above, but 5 mM okadaic acid was added to inhibit the activity of PP1 and PP2A family Ser/Thr-specific phosphoprotein phosphatases²⁵.

In vitro kinase assays

The recombinant protein was expressed in Rosetta (DE3) and affinity purified according to the instructions described above. Two micrograms of GST or GST-FL7 proteins were incubated with 0.5 µg of MBP-ABI1 with or without 0.5 µM ABA in 20 µL of phosphatase buffer (50 mM Tris-HCl (pH 7.5), 10 mM MgCl₂, 1 mM EGTA and 0.1% [v/v] β-mercaptoethanol). After 15 min of incubation, 1 µg of SnRK2.6-HIS protein was added to the reaction buffer and incubated for another 15 min. Then, the reaction buffer was mixed with 1 µM ATP and phosphatase inhibitor cocktail 3 to a total volume of 25 µL and incubated for 30 min at 25 °C. The reactions were stopped by the addition of SDS loading buffer, and the mixtures were separated by SDS–PAGE. The anti-phospho-S175-SnRK2 antibody was previously reported (1:5000)⁵². The protein level was analysed by Coomassie Brilliant Blue (CBB) staining.

SnRK2.6 kinase activity in vivo

To assess the kinase activity of snRK2.6 in vivo, total protein was extracted from Col-0, fl7 and OE-FL7 with lysis buffer (100 mM HEPES (pH 7.8), 10 mM Na₃VO₄, 10 mM NaF, 5 mM EDTA, 5 mM EGTA, 10 mM DTT, 1 mM PMSF, and 5% glycerin and protease inhibitor cocktail), separated by SDS–PAGE and detected by immunoblotting with anti-phosphorylation antibodies against Ser175 in SnRK2.6³⁵.

Arabidopsis protoplast preparation and transfection

Fully expanded rosette leaves from 4-week-old Col-0 plants were enzymatically digested to release protoplasts⁵⁵, which were washed and resuspended at ~2 × 10⁶ cells/ml. For each transfection, 100 μl protoplasts (~2 × 10⁵ cells) were mixed with 20 μg plasmid DNA and an equal volume of PEG solution (40% PEG4000, 0.2 M mannitol, 100 mM CaCl₂), incubated 10–15 min, then diluted with W5 (154 mM NaCl, 125 mM CaCl₂, 5 mM KCl, and 2 mM MES; pH 5.7) and recovered. Transfected protoplasts were incubated in the dark at 22 °C for 12–24 h before downstream assays.

Phylogenetic analysis

Multiple alignments of full-length aa sequences were aligned using MUSCLE. Phylogenetic analysis was performed using full-length sequences with the MEGA X program by the maximum-likelihood method with 100 bootstrap samples and the following parameters: Poisson model, uniform rates, and complete deletion.

RNA-seq analysis

For RNA sequencing, total RNA from nine-day-old seedlings treated with or without 300 mM mannitol on 1/2 MS solid medium for 12 h³⁵ was extracted using an RNA-Simple Total RNA Kit (Zoman Biotech) and prepared for RNA sequencing at Novogene (Beijing, China). The sequencing data were initially filtered, after which the clean reads were mapped to the Arabidopsis genome using HISAT2⁵⁷. The transcripts per million (TPM) value of each gene was quantified using the Stringtie tool⁵⁷. DEGSeq2 facilitated the identification of differentially expressed genes (DEGs) across samples, focusing on genes that exhibited a |log2(fold change)| ≥ 1 and a P ≤ 0.05 as DEGs⁵⁸. Gene expression patterns were clustered using the pheatmap package in R, employing the ward.D2 clustering algorithm. GO enrichment analyses were conducted with the GO.db package in R.

Statistical analysis

All data are shown as the means ± standard deviations (SD) from at least three biological replicates or from three technical replicates in one of three experiments with similar results. Two-tailed Student’s t test was used to compare the means between two samples (* and ** represent P < 0.05 and 0.01, respectively). One- or two-way analysis of variance (ANOVA) was used to test the significance of the difference among different group means (different lowercase letters indicate significant differences; P < 0.05). Detailed statistical reports are provided in the source data for the figures.

Accession numbers

Sequence data from this article can be found in the TAIR (The Arabidopsis Information Resource) databases under the following accession numbers: FL7 (At4g16670), ABI1 (At4g26080), ABI2 (At5g57050), HAB1 (At1g72770), HAB2 (At1g17550), HAI1 (At5g59220), HAI2 (At1g07430), HAI3 (At2g29380), AHG1 (At5g51760), AHG3 (At3g11410), PP2C-D (At5g02760), PYL10 (At4g27920), SnRK2.6 (At4g33950), MPK6 (At2g43790), MKK4 (At1g51660), RD29B (At5g52300), AFP1 (At1g69260), AtNAP (At1g69490), P5CS1 (At2g39800), SAG13 (At2g29350), ABF2 (At1g45249), COR413IM1 (At1g29395), EGY3 (At1g17870), GOLS2 (At1g56600), LEA7 (At1g52690), PNP-A (At2g18660), RAB18 (At5g66400), SINA2 (At3g13672) and ABA1 (At5g67030).

Reporting summary

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

Data availability

Source data and uncropped western blot membranes are provided with this paper. The RNA-seq raw read data generated in this study have been deposited in the National Genomics Data Center database under accession code CRA020685. Source data are provided with this paper.

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Acknowledgements

The anti-phosphorylation antibodies for Ser175 in SnRK2.6 is a gift from Y.Z. in CAS Center for Excellence in Molecular Plant Sciences, China. We thank Pro. Assaf Mosquna in the Hebrew University of Jerusalem, Israel, and Pro. Yajin Ye in Nanjing Forestry University, China for helpful suggestions and for manuscript editing. The work was supported by the Natural Science Foundation of Jiangsu Province, China (BK20230984, G.A.), the National Natural Science Foundation of China (32402315, G.A.; 32230089, D.D.; 32372493, D.S.), the China Agriculture Research System (CARS-21, D.D.) and the National Key Research and Development Program (2024YFD1401103, G.A.).

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These authors contributed equally: Tianli Li, Zitong Yang.

Authors and Affiliations

State Key Laboratory of Agricultural and Forestry Biosecurity, Nanjing Agricultural University, Nanjing, China
Tianli Li, Zitong Yang, Yuke Li, Xianglan Li, Xu Lu, Xinyi Gu, Daolong Dou, Danyu Shen & Gan Ai
College of Plant Protection, Nanjing Agricultural University, Nanjing, China
Tianli Li, Zitong Yang, Yuke Li, Xianglan Li, Xu Lu, Xinyi Gu, Daolong Dou, Danyu Shen & Gan Ai
CAS Center for Excellence in Molecular Plant Sciences, The Chinese Academy of Sciences, Shanghai, China
Guojun Li & Yang Zhao
Key Laboratory of Plant Carbon Capture, The Chinese Academy of Sciences, Shanghai, China
Guojun Li & Yang Zhao
University of Chinese Academy of Sciences, Beijing, China
Guojun Li & Yang Zhao
Department of Applied Bioinformatics, University of Goettingen, Goettingen, Germany
Jan de Vries
Campus Institute Data Science (CIDAS), University of Goettingen, Goettingen, Germany
Jan de Vries
Department of Applied Bioinformatics, Goettingen Center for Molecular Biosciences (GZMB), University of Goettingen, Goettingen, Germany
Jan de Vries
Guangxi Key Laboratory of Agric-Environment and Agric-Products Safety, College of Agriculture, Guangxi University, Nanning, Guangxi, China
Zhaojie Chen
Academy for Advanced Interdisciplinary Studies, Nanjing Agricultural University, Nanjing, China
Daolong Dou

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Tianli Li
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Zitong Yang
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Guojun Li
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Jan de Vries
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Yuke Li
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Xianglan Li
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Xu Lu
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Xinyi Gu
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Yang Zhao
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Daolong Dou
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Danyu Shen
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Contributions

G.A., D.S., and D.D. conceived and designed the project. T.L., Z.Y., G.L., Y.L., X.Li., X.Lu., X.G., and Z.C. performed experiments and analyzed data. D.S., J.d.V., Y.Z., G.A., and D.D. wrote the manuscript. All authors reviewed and approved the final manuscript.

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Correspondence to Danyu Shen or Gan Ai.

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Nature Communications thanks Yoichi Sakata, and Cun Wang for their contribution to the peer review of this work. A peer review file is available.

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Li, T., Yang, Z., Li, G. et al. FL7 is an ancient ABA-independent inhibitor of PP2C-As regulating plant stress responses. Nat Commun 16, 11119 (2025). https://doi.org/10.1038/s41467-025-66086-z

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Received: 05 June 2024
Accepted: 30 October 2025
Published: 15 December 2025
Version of record: 15 December 2025
DOI: https://doi.org/10.1038/s41467-025-66086-z