SlMED25‐SlPHR3‐SlSPX2 module fine‐tunes SlPHR3‐mediated transcriptional activation of phosphate starvation response in tomato

ABSTRACT

Phosphate starvation response (PHR) transcription factors are master regulators of the plant phosphate (Pi) starvation response (PSR), yet the mechanisms governing the dynamic control of their transcriptional activity remain elusive. Here, we report a dual regulatory module comprising the coactivator SlMED25 and the corepressor SlSPX2 that fine‐tunes SlPHR3 activity in Solanum lycopersicum (tomato). Genetic and biochemical evidence collectively confirmed that SlPHR3 acts as the central regulator orchestrating tomato PSR. Specifically, the mediator subunit SlMED25 interacts with the N‐terminal domain (NTD) of SlPHR3 to recruit RNA polymerase II (Pol II) to SlPHR3 target promoters in a SlPHR3‐dependent fashion, whereas SlSPX2 binds to the same NTD of SlPHR3 to robustly suppress its transcriptional activity. Biochemical assays further demonstrated that SlSPX2 and SlMED25 compete for binding to SlPHR3, with SlSPX2 exhibiting higher binding affinity. This competitive binding module functions as a key molecular switch that mediates dynamic PSR modulation in tomato, thus yielding distinct functional outputs in response to varying intracellular Pi levels. Our findings uncover a previously uncharacterized regulatory layer in the PSR network, wherein a Mediator subunit and a Pi‐sensing protein modulate PHR activity via competitive binding, thereby enhancing the mechanistic insight into Pi homeostasis regulation in plants.

The mediator subunit SlMED25 and the co‐repressor SlSPX2 competitively bind the N‐terminal domain of the phosphate starvation response transcription factor SlPHR3 in tomato to form a sensitive molecular switch, which dynamically modulates phosphate starvation responses and maintains phosphate homeostasis, offering valuable targets for breeding low phosphorus tolerant crops.

INTRODUCTION

Phosphorus (P) is one of the essential macronutrients for plant growth and development (Paz‐Ares et al., 2022; Puga et al., 2024; Yang et al., 2024). Plants primarily absorb and utilize P in the form of inorganic phosphate (Pi), which is readily immobilized by metal ions, thereby severely limiting the effective availability of P. Although total P content in soil is substantial, the majority exists as organic phosphorus or insoluble phosphate (Shen et al., 2011). To mitigate this limitation, excessive phosphate fertilizer is commonly applied to enhance crop yield (Shen et al., 2011). However, phosphate rock resources are finite and non‐renewable, posing a severe constraint on the sustainable development of modern agriculture (Neset and Cordell, 2012; Xiao et al., 2022). Furthermore, excessive application of Pi fertilizers can trigger adverse environmental consequences such as water eutrophication (Conley et al., 2009; Lopez‐Arredondo et al., 2014). Faced with the scarcity of phosphate rock resources and low P use efficiency, improving crop P absorption and utilization efficiency has emerged as a critical and pressing challenge in agricultural production (Vance et al., 2003; Heuer et al., 2017).

The MYB‐CC transcription factors PHR‐mediated transcriptional reprogramming serve as the central hub in phosphate starvation response (PSR) (Bustos et al., 2010; Wu et al., 2013; Sega and Pacak, 2019; Paz‐Ares et al., 2022; Yang et al., 2024). In Arabidopsis, AtPHR1 regulates the expression of downstream PSR genes by binding to the PHR1 binding site (P1BS) element, thereby participating in PSR (Rubio et al., 2001; Sun et al., 2016). The homologs of AtPHR1, OsPHR2 in rice, SlPHR3 and SlPHR4 in tomato, ZmPHR1 in maize, PvPHR1 in common bean, and BnPHR1 in Brassica napus were also served as the central players in regulating PSR (Zhou et al., 2008; Ren et al., 2012; Wu et al., 2023; Tian et al., 2024; Lin et al., 2025). These studies indicate that the functions of the PHR proteins are highly conserved across different species. AtPHR1 and AtPHL2 play a key role in remodeling chromatin accessibility during PSR (Barragan‐Rosillo et al., 2021). The PHR signaling pathway is also involved in Pi uptake from the soil through mycorrhizal symbiosis (Shi et al., 2021; Liao et al., 2022). Additionally, AtPHR1‐mediated PSR suppresses plant immune responses under Pi starvation in Arabidopsis (Castrillo et al., 2017; Motte and Beeckman, 2017; Tang et al., 2022; Inoue et al., 2024). ZmPHR1 positively regulates drought resistance by modulating PSR in maize (Tian et al., 2024). PSR‐specific mobile mRNAs are transported to recipient tissues to modulate PSR, and the mobility of these mRNAs is largely dependent on PHR function (Dong et al., 2025). In response to fluctuations in available Pi levels within plants, the post‐transcriptional modification of PHR1 is finely regulated. In Arabidopsis and rice, the protein abundance of AtPHR1 and OsPHR2 is controlled by 26S proteasome‐mediated degradation (Guo et al., 2022; Park et al., 2023). In rice, GSK3/SHAGGY‐LIKE KINASE2 (GSK2), a core inhibitor of the brassinosteroid (BR) signaling pathway, directly phosphorylates OsPHR2, thereby suppressing its DNA‐binding activity (Zhang et al., 2024).

Direct transcriptional targets of PHR1 in Arabidopsis can be categorized into five distinct classes: (i) SPX1 and SPX2; (ii) the Pi transporter PHT1; (iii) miR399 and miR827, which specifically and respectively target the mRNAs of PHO2 (an E2 ubiquitin conjugase) and NLA (an E3 ubiquitin ligase); (iv) the long non‐coding RNAs (lncRNAs) IPS1/At4, which sequester miR399; and (v) strigolactone (SL) biosynthesis enzymes (SLBs). PHO2 and NLA mediate the degradation of PHT1 and/or PHO1 (a Pi transporter involved in xylem Pi loading in roots) via the ubiquitin‐proteasome pathway (Paz‐Ares et al., 2022). Inositol pyrophosphate (InsPs), ATP, and Pi are the major forms of phosphorus in plants (Wild et al., 2016). Under Pi starvation, the levels of ATP and Pi in plants decrease, while the synthesis of InsP₈ depends on ATP and Pi. Thus, the intracellular InsP₈ level in plants serves as an intracellular Pi signal in plants (Dong et al., 2019; Zhu et al., 2019). SPX (Syg1/Pho81/XPR1) proteins serve as the repressors of PHR proteins (Zhou et al., 2021; Yang et al., 2024), and also Pi sensors by binding InsP₈ with high affinity (Puga et al., 2014; Wang et al., 2014). Through structure‐function analyses of AtPHR1 in Arabidopsis and OsPHR2 in rice, the mechanism by which SPX proteins inhibit PHR's function in the presence of InsP₈ has been elucidated (Ried et al., 2021; Zhou et al., 2021). Under Pi‐sufficient conditions, InsP8 binding to SPX1, SPX2, and SPX4 stabilizes the interaction between PHR1 and SPXs, disrupting PHR1 dimerization, blocking PHR1's DNA‐binding activity, and sequestering PHR1 in the cytoplasm. In the Pi‐deplete condition, InsP₈ levels decrease, and PHR1 is released from the PHR1‐SPX complex, activating PSR (Paz‐Ares et al., 2022).

The Mediator complex is a eukaryotic multi‐subunit transcriptional co‐activator that acts as a bridge between signal‐responsive transcription factors (TFs) and the core RNA polymerase II (Pol II) transcriptional machinery, as well as several other transcription‐associated factors. Thus, Mediator functions as a central integrative hub in regulating multiple aspects of gene transcription (Zhai et al., 2020; Freytes et al., 2024; Wu et al., 2025). First isolated from Arabidopsis thaliana, the plant Mediator complex has been shown to comprise 29 conserved subunits and six plant‐specific subunits (Bourbon, 2008; Mathur et al., 2011). Although several plant Mediator subunits have been implicated in the regulation of plant development and adaptive responses (Chen et al., 2012; Yang et al., 2016; Chen et al., 2022; Freytes et al., 2024), the molecular underpinnings of how plant Mediator modulates PSR remain incomplete. Specifically, the Mediator subunit(s) that modulate SlPHR‐mediated transcriptional regulation of PSR in tomato remain elusive.

Here, we report the functional role of Mediator subunit SlMED25 in regulating PSR in tomato. We demonstrate that SlMED25 physically interacts with SlPHR3, the key transcription factor governing PSR in tomato. Notably, SlSPX2 and SlMED25 engage in a competitive binding event for SlPHR3, with SlSPX2 exhibiting a higher binding affinity for SlPHR3 than SlMED25. This competitive binding module functions as a key molecular switch for dynamic PSR modulation in tomato, yielding divergent functional outcomes under varying intracellular Pi conditions.

RESULTS

SlPHR3 is the key transcription factor mediating tomato PSR

Phylogenetic analysis demonstrated that tomato SlPHR3 (Solyc09g072830) exhibits high homology to Arabidopsis AtPHR1 (Figure S1). To further dissect the specific function of SlPHR3 in the transcriptional regulation of PSR, loss‐of‐function mutants of SlPHR3 were generated via the CRISPR/Cas9 gene‐editing system. Sequence alignment revealed a 1‐bp insertion in slphr3‐22 and a 7‐bp deletion in slphr3‐210, both within the open reading frame (ORF), leading to frameshift mutations and the formation of premature stop codons (TAG or TGA) (Figure S2). Given that Pi deficiency (–Pi) typically inhibits plant growth and decreases internal Pi levels, the inhibition rates of shoot height, fresh weight, and Pi content were employed as indicators of tolerance to –Pi. The results demonstrated that the inhibition rates of these three traits in slphr3‐22 and slphr3‐210 were significantly higher than those in WT (Figure 1A–D). These findings indicate that SlPHR3 positively regulates PSR in tomato.

Figure 1.

SlPHR3 positively regulates phosphate starvation response (PSR) in tomato

(A) Seedlings of wild‐type (WT) and slphr3 grown under +Pi (1 mM KH₂PO₄) or −Pi (0.05 mM KH₂PO₄) for 21 d. Scale bars = 10 cm. (B–D) Inhibition rate of shoot height (B), fresh weight (C), and Pi content (D) in WT and slphr3 seedlings under +Pi and −Pi. Data are means ± SD (n ≥ 5 plants). (E–H) RT‐qPCR analysis of SlSPX1 (E), SlSPX2 (F), SlPAP26 (G), and SlPT1 (H) in WT and slphr3 under +Pi or −Pi. Data represent means ± SD of three independent experiments (n = 3). (I, J) SlSPX1 (I) and SlPAP26 (J) promoter‐binding patterns of SlPHR3. Vertical black lines indicate putative PHR1 binding site (P1BS); short red lines indicate fragments used for ChIP‐qPCR; red arrowheads indicate DNA probes used in EMSAs. (K, L) SlPHR3 enrichment at the P1BS motif in SlSPX1 (K) and SlPAP26 (L) promoters in 35S::SlPHR3‐GFP under +Pi and −Pi by ChIP‐qPCR. SlACTIN2 was used as a negative control. Data represent means ± SD of three independent experiments (n = 3). (M, N) EMSA showing the binding of SlPHR3‐GST to the SlSPX1 (M) and SlPAP26 (N) promoters. GST alone and mutated probes were used as negative controls. Molar excesses (5‐ and 10‐fold) of unlabeled or mutated probes were used as competitors. In the mutant competitor (Mu‐competitor), the P1BS motif was mutated to a poly‐A sequence. Experiments were repeated at least three times independently with similar results. Asterisks indicate significant differences (ns: no significant difference, *P < 0.05, **P < 0.01, ****P < 0.0001, Student's t‐tests) in (B–D, K, L). (E–H) Different lowercase letters indicate significant differences, as determined by Two‐way ANOVA with Tukey's test, P ＜ 0.05.

To clarify the biological function of SlPHR3 in PSR of tomato, we generated SlPHR3 overexpression lines (35S::SlPHR3‐GFP), designated as SlPHR3‐OE‐1 and SlPHR3‐OE‐3. RT‐qPCR validation demonstrated that SlPHR3 transcript abundances in SlPHR3‐OE‐1 and SlPHR3‐OE‐3 were significantly elevated compared to WT (Figure S3B). Under –Pi condition, SlPHR3‐OE plants exhibited significantly alleviated growth inhibition and improved Pi homeostasis relative to WT (Figures S3A, C–E). These results further validate that SlPHR3 functions as a positive regulator of PSR in tomato.

To further investigate the regulatory role of SlPHR3 in PSR gene expression, we determined the transcript abundances of four key PSR‐related genes (SlSPX1, SlSPX2, SlPAP26, and SlPT1) in WT and slphr3 under Pi‐sufficient (+Pi) and –Pi conditions. Under +Pi conditions, no significant differences in the expression levels of these genes were detected across the genotypes. In contrast, under –Pi conditions, the transcript abundances of these PSR genes were significantly lower in slphr3 compared to those in WT (Figure 1E–H). Conversely, SlPHR3‐OE lines showed significantly elevated expression levels of these PSR genes relative to WT under –Pi conditions (Figure S3F–I). Collectively, these findings demonstrate that SlPHR3 positively regulates the expression of key PSR genes in response to –Pi.

To investigate whether SlPHR3 directly regulates SlSPX1 and SlPAP26 in tomato through binding to their promoter regions, chromatin immunoprecipitation (ChIP) assays were performed using SlPHR3‐OE lines. Since AtPHR1 specifically binds to the P1BS motif in the promoters of its target genes (Wang et al., 2018), we analyzed the promoter regions of SlSPX1 and SlPAP26 and identified four potential P1BS motifs (Figure 1I, J). ChIP‐qPCR assays revealed that SlPHR3 was specifically enriched at the P1BS motif‐containing regions within the SlSPX1 and SlPAP26 promoters under –Pi conditions (Figure 1K, L). Furthermore, electrophoretic mobility shift assays (EMSAs) demonstrated that a recombinant SlPHR3‐GST (glutathione S‐transferase) fusion protein binds to the promoter regions of SlSPX1 and SlPAP26 in a P1BS‐dependent manner (Figure 1M, N). These findings confirm that SlPHR3 positively regulates the expression of SlSPX1 and SlPAP26 under –Pi conditions by directly binding to their promoters.

Taken together, our findings establish that SlPHR3 positively regulates PSR in tomato by directly modulating the expression of core PSR‐related genes.

SlMED25 acts as a coactivator of SlPHR3 during PSR in tomato

Given that the Mediator complex serves as a critical bridge between transcription factors and RNA polymerase II to modulate gene transcription, we set out to identify the specific Mediator subunit(s) required for transducing the transcriptional activation function of SlPHR3 during PSR in tomato. Our previous reports have demonstrated that the tomato Mediator subunit SlMED25 regulates hypocotyl elongation and fruit ripening (Sun et al., 2020; Deng et al., 2023). Based on this, we postulated that SlMED25 contributes to SlPHR3‐mediated PSR. We analyzed the phenotypic characteristics of SlMED25‐antisense (SlMED25‐AS) lines under –Pi, in which SlMED25 transcript levels are significantly downregulated relative to WT (Liu et al., 2019). Under –Pi, SlMED25‐AS plants displayed significantly higher inhibition rates of shoot height, fresh weight, and Pi content compared to WT (Figure 2A–D). Consistent with the phenotypic observations, –Pi induced expression of core PSR‐related genes (SlSPX1, SlSPX2, SlPAP26, and SlPT1) was markedly downregulated in SlMED25‐AS lines relative to WT (Figure 2E–H). These findings demonstrate that SlMED25 positively regulates PSR in tomato.

Figure 2.

SlMED25 acts as a coactivator of SlPHR3 to positively regulate PSR

(A) Seedlings of WT and SlMED25‐AS under +Pi or −Pi. Scale bar = 10 cm. (B–D) Inhibition rate of shoot height (B), fresh weight (C), and Pi content (D) in WT and SlMED25‐AS seedlings under +Pi and −Pi. Data are means ± SD (n ≥ 5 plants). (E–H) RT‐qPCR analysis of SlSPX1 (E), SlSPX2 (F), SlPAP26 (G), and SlPT1 (H) in WT and SlMED25‐AS seedlings under +Pi or −Pi. Data represent means ± SD of three independent experiments (n = 3). (I) Schematic diagram of effector constructs and LUC reporter construct used in transient expression assays in (J). (J) Quantification of luciferase (LUC) luminescence intensity in Nicotiana benthamiana leaves at 60 h post‐coinfiltration with the proSlPAP26::LUC reporter construct and effector constructs, with infiltration of the proSlPAP26::LUC reporter construct alone as the control. Data are means ± SD (n = 5 leaves). Asterisks indicate significant differences (*P < 0.05, ***P < 0.001, ****P < 0.0001, Student's t‐test) in (B–D). (E–H, J) Different lowercase letters indicate significant differences, as determined by two‐way ANOVA with Tukey's test, P ＜ 0.05.

Building on these observations, we next investigated whether SlMED25 functions as a transcriptional coactivator of SlPHR3. To address this question, we generated a luciferase (LUC) reporter construct controlled by the 2,177‐bp SlPAP26 promoter (proSlPAP26::LUC). Transient co‐expression of SlPHR3 with this reporter construct in Nicotiana benthamiana leaves led to a significant increase in LUC activity, confirming that SlPHR3 directly activates SlPAP26 expression. Notably, co‐expression of SlMED25 further potentiated SlPHR3‐dependent transactivation of the reporter. In contrast, SlMED25 alone had no significant effect on reporter activity (Figure 2I, J). Based on the above results, we found that the positive regulation of PSR genes by SlMED25 is dependent on the presence of SlPHR3. To verify this conclusion, we conducted RT‐qPCR analysis to detect the expression changes of PSR genes following SlMED25 overexpression in WT (SlMED25‐OE) and slphr3 (SlMED25‐OE/slphr3) backgrounds (Figure S4). The results showed that under –Pi, SlMED25‐OE significantly upregulated the expression of SlSPX1 and SlPAP26. Notably, the promotion of SlSPX1 and SlPAP26 expression was completely abolished in SlMED25‐OE/slphr3. This further confirms that SlMED25 cannot function independently of SlPHR3; instead, it acts as a transcriptional co‐activator of SlPHR3 to exert its regulatory roles.

These data substantiate that SlMED25 acts as a transcriptional coactivator of SlPHR3 to modulate PSR in tomato.

SlMED25 interacts with SlPHR3 to positively modulate PSR in tomato

The identification of SlMED25 as a coactivator of SlPHR3 prompted us to hypothesize that these two proteins may engage in a direct physical interaction. To test this, we first performed Y2H assays by fusing full‐length SlPHR3 to the GAL4 activation domain (pGADT7, AD) and SlMED25 to the DNA‐binding domain (pGBKT7, BD). Results confirmed a direct interaction between SlPHR3 and SlMED25 in yeast (Figure 3A). We next conducted in vitro pull‐down assays using purified recombinant proteins: Maltose‐binding protein (MBP)‐tagged SlMED25, but not MBP alone, specifically pulled down GST‐tagged SlPHR3 (Figure 3B), verifying their direct physical interaction in vitro.

Figure 3.

SlMED25 physically interacts with SlPHR3

(A) Y2H assays showing interactions of SlPHR3 with SlMED25. SlMED25 was fused with the DNA‐binding domain (BD) in pGBKT7, and SlPHR3 was fused with the activation domain (AD) in pGADT7. Yeast transformants were grown on SD medium lacking Leu and Trp (SD/−2) as a transformation control or SD medium lacking Ade, His, Leu, and Trp (SD/−4) to test protein–protein interactions. The empty pGADT7 or pGBKT7 vectors were co‐transformed with SlMED25‐BD or SlPHR3‐AD in parallel as negative controls. (B) In vitro pull‐down assays to verify the interaction between SlMED25 and SlPHR3. Purified SlPHR3‐GST was incubated with MBP or SlMED25‐MBP, and proteins recovered from the MBP pull‐down assays were detected by immunoblotting using anti‐MBP and anti‐GST antibodies. Asterisks indicate the specific bands of recombinant proteins. (C) Co‐IP assays to verify in vivo interactions between SlPHR3 and SlMED25. The indicated construct combination was co‐infiltrated into Nicotiana benthamiana leaves, and then the protein extracts were immunoprecipitated with GFP antibody‐bound agarose beads. Asterisks indicate the specific bands of recombinant proteins. (D) In vivo LCI assays showing the association between SlMED25 and SlPHR3. Various constructs were co‐infiltrated into N. benthamiana leaves, and LUC activity was captured (upper panel). Different construct combinations co‐infiltrated into N. benthamiana leaves are shown in the lower four squares. The pseudocolor bar shows the range of luminescence intensity. (E, F) Mapping of SlPHR3 and SlMED25 domains involved in the SlPHR3‐SlMED25 interaction by Y2H. SlMED25 and its derivatives were cloned into pGBKT7 (E), and SlPHR3 and its derivatives cloned into pGADT7 (F) were co‐transformed into yeast. The yeast transformants were grown on SD/− 2 or on SD/− 4. The left panel indicates different truncations of SlMED25 and SlPHR3. Experiments in (A–F) were repeated at least three times independently with similar results.

To further validate this interaction in planta, we transiently co‐expressed SlMED25‐myc and SlPHR3‐GFP in N. benthamiana leaves and performed Co‐IP assays. Protein extracts were immunoprecipitated using an anti‐GFP antibody, and the immunoprecipitates were detected by immunoblotting with an anti‐myc antibody. SlMED25‐myc was specifically detected in samples co‐expressing SlPHR3‐GFP, but not in those expressing GFP alone (Figure 3C). Additionally, we employed a luciferase complementation imaging (LCI) assay in N. benthamiana: Strong luminescence signals were detected in leaves co‐expressing cLUC‐SlMED25 and SlPHR3‐nLUC, whereas no detectable signals were observed in control combinations (cLUC‐SlMED25 + SCaBP8‐nLUC or cLUC‐SlULP1 + SlPHR3‐nLUC) (Figure 3D).

To map the SlMED25‐interacting domain(s) within SlPHR3, we generated a series of truncated SlPHR3 constructs (full‐length and various deletion mutants) (Figure 3E) fused to the AD vector, and co‐transformed them with SlMED25‐BD in yeast. Y2H assays showed that the N‐terminal domain (NTD) of SlPHR3 is responsible for mediating the interaction with SlMED25. Deletion of the NTD completely abrogated the interaction, whereas deletion of the MYB domain or coiled‐coil (CC) domain had no significant impact (Figure 3E). These results indicate that the NTD of SlPHR3 is both necessary and sufficient for binding to SlMED25.

Similarly, to define the SlPHR3‐interacting domain in SlMED25, we divided the protein into four functional segments based on previous reports (Chen et al., 2012; Sun et al., 2020): the von Willebrand Factor A (vWF‐A) domain, a nonconserved middle domain (MD), the ACID domain, and a glutamine (Gln)‐rich domain (GD). Each segment, together with full‐length SlMED25, was fused to the BD vector and co‐expressed with full‐length SlPHR3 fused with AD in yeast. Y2H results confirmed that the ACID domain of SlMED25 is both essential and sufficient for its interaction with SlPHR3 (Figure 3F).

Collectively, these findings provide compelling evidence that SlMED25 physically interacts with SlPHR3 both in vitro and in vivo.

SlMED25 recruits Pol II to the promoters of SlPHR3 target genes

It is well established that transcription factors recruit MED25 to the promoter regions of their target genes to activate transcription (Chen et al., 2012; Liu et al., 2019; Sun et al., 2020; Deng et al., 2023). To evaluate the enrichment of SlMED25 at the chromatin regions of SlPHR3 target genes, we performed ChIP‐qPCR assays using 35S::SlMED25‐GFP transgenic plants (Liu et al., 2019). Given that SlMED25 is ubiquitously expressed in various tissues of WT plants (Figure S5), we analyzed 6 d‐old whole seedlings of SlMED25‐GFP plants. ChIP‐qPCR results revealed significant enrichment of SlMED25 at the P1BS motif‐containing regions within the SlSPX1 and SlPAP26 promoters under –Pi (Figure 4A, B). To determine whether SlPHR3 is required for this SlMED25 enrichment, we introgressed the SlMED25‐GFP transgenic plants into slphr3‐22 via genetic crossing. ChIP‐qPCR analysis showed that –Pi‐induced enrichment of SlMED25 at the SlSPX1 and SlPAP26 promoters was substantially attenuated in slphr3‐22 compared to WT (Figure 4A, B). These findings demonstrate that the recruitment of SlMED25 to SlPHR3 target promoters and its subsequent functional activity are dependent on SlPHR3.

Figure 4.

SlMED25 recruits RNA polymerase II (Pol II) to the promoters of SlPHR3 target genes

(A) SlSPX1 and SlPAP26 promoter‐binding patterns of SlPHR3. Vertical black lines indicate putative P1BS; short red lines indicate fragments used for ChIP‐qPCR. (B, C) ChIP‐qPCR assays showing the enrichment of SlMED25 (B) and Pol II (C) on SlSPX1 (left) and SlPAP26 (right) promoters under +Pi and −Pi. SlACTIN2 was used as a negative control. Data represent means ± SD of three independent experiments (n = 3). Asterisks indicate significant differences (****P < 0.0001, Student's t‐test) in (B, C).

Previous reports have demonstrated that SlMED25 can physically associate with both DNA‐bound transcription factors and RNA polymerase II (Pol II), thereby acting as a molecular bridge to facilitate transcriptional activation (Chen et al., 2012). In this context, our observation that SlMED25 interacts with SlPHR3 and modulates its transcriptional activity suggests that loss of SlMED25 function may compromise the recruitment of the Pol II transcriptional machinery during –Pi‐induced PSR gene transcription. To test this, we examined whether SlMED25 depletion affects the recruitment of the C‐terminal domain (CTD) of Pol II's largest subunit to SlPHR3 target promoters. ChIP‐qPCR assays confirmed that –Pi‐induced recruitment of the Pol II CTD to the SlSPX1 and SlPAP26 promoters was markedly reduced in SlMED25‐AS plants relative to WT (Figure 4A, C).

Collectively, these findings provide compelling evidence that SlMED25 facilitates the recruitment of Pol II to the promoters of SlPHR3 target genes, thereby establishing a functional connection between SlPHR3 and the general transcriptional machinery.

SlSPX2 interacts with the NTD domain of SlPHR3

Our findings establish SlMED25 as a transcriptional coactivator of SlPHR3. Under –Pi conditions, SlMED25 contributes to PSR activation, whereas under +Pi conditions, transcriptional repression of SlPHR3 activity is required. Prior studies have confirmed SPX proteins as Pi sensors: Under +Pi conditions, their SPX domains bind InsP8 to either suppress PHR1 transcriptional activity or attenuate PHR1 through cytoplasmic sequestration (Zhu et al., 2019). The tomato genome harbors five putative SPX proteins, designated SlSPX1–SlSPX5. Phylogenetic analysis further revealed that SlSPX1, SlSPX2, and SlSPX3 are orthologous to Arabidopsis AtSPX1, AtSPX2, and AtSPX3, respectively, whereas SlSPX4 and SlSPX5 both exhibit sequence homology to Arabidopsis AtSPX4 (Figure S6). To investigate the subcellular localization of tomato SlSPX proteins, we generated 35S::SlSPX1‐GFP, 35S::SlSPX2‐GFP, 35S::SlSPX3‐GFP, 35S::SlSPX4‐GFP, and 35S::SlSPX5‐GFP constructs and transiently expressed them in N. benthamiana leaves. DAPI staining confirmed that the GFP signals of SlSPX1, SlSPX2, and SlSPX3 overlapped with nuclear DAPI signals (Figure 5A), indicating their nuclear localization. In contrast, SlSPX4 and SlSPX5 localized to the cytoplasm and plasma membrane, respectively (Figure S7).

Figure 5.

SlSPX2 physically interacts with SlPHR3

(A) Nuclear localization of SlSPX1‐GFP, SlSPX2‐GFP, and SlSPX3‐GFP by transient expression in Nicotiana benthamiana leaves. Nuclei were counterstained with DAPI (4′,6‐Diamidino‐2‐Phenylindole). Scale bars = 20 μm. (B) Y2H assays showing that SlPHR3 interacts with SlSPX1, SlSPX2, and SlSPX3. SlSPX1, SlSPX2, and SlSPX3 were fused with BD in pGBKT7, and SlPHR3 was fused with AD in pGADT7. Transformed yeast was grown on selective SD/−2, SD/−3, or SD/−4 medium to assess protein‐protein interaction. As negative controls, the empty pGADT7 or pGBKT7 vector was co‐transformed with SlSPX1‐BD, SlSPX2‐BD, SlSPX3‐BD, or SlPHR3‐AD in parallel experiments. (C) In vitro pull‐down assays to verify SlSPX2 and SlPHR3 interaction. Proteins were pulled down by Amylose Resin, and the eluates were analyzed by immunoblotting with the indicated antibodies. Asterisks indicate the specific bands of recombinant proteins. (D) Co‐IP assays to verify in vivo interaction of SlPHR3 and SlSPX2. The indicated construct combination was co‐infiltrated into N. benthamiana leaves, and then the protein extracts were immunoprecipitated with GFP antibody‐bound agarose beads. Asterisks indicate the specific bands of recombinant proteins. (E) In vivo LCI assays showing the association of SlSPX2 and SlPHR3. Various constructs were co‐infiltrated into N. benthamiana leaves, and LUC activity was captured (upper panel). Different construct combinations co‐infiltrated into N. benthamiana leaves are shown in the lower four squares. The pseudocolor bar shows the range of luminescence intensity. (F) Mapping of SlPHR3 domains involved in SlPHR3‐SlSPX2 interaction by Y2H. SlPHR3 and its derivatives were cloned into pGADT7, and SlSPX2 was cloned into pGBKT7. The transformed yeast was grown on SD/−2/−3/−4. The left panel indicates different truncations of SlPHR3. Experiments were repeated at least three times independently with similar results in (A–F).

Given that SPX proteins repress PHR1 function and SlSPX1–SlSPX3 are nuclear‐localized, we hypothesized that these three SlSPXs may interact directly with SlPHR3. Y2H assays—using full‐length SlSPX1–SlSPX3 fused to the BD domain and SlPHR3 fused to the AD domain—revealed that all three SlSPXs interact with SlPHR3 (Figure 5B). Among them, SlSPX2 exhibited the strongest interaction, enabling robust yeast growth on selective medium, whereas SlSPX1 and SlSPX3 showed weaker binding. We therefore focused subsequent analyses on SlSPX2. In vitro pull‐down assays using purified recombinant proteins demonstrated that SlPHR3‐MBP, but not MBP alone, specifically pulled down SlSPX2‐GST (Figure 5C), confirming their direct physical interaction. Co‐IP assays in N. benthamiana leaves further validated this interaction in planta: SlSPX2‐Flag was co‐immunoprecipitated with SlPHR3‐GFP, but not with GFP alone (Figure 5D). Additionally, LCI assays showed strong luminescence when cLUC‐SlSPX2 and SlPHR3‐nLUC were co‐expressed, while control combinations yielded no detectable signal (Figure 5E). Collectively, these data confirm that SlSPX2 interacts with SlPHR3 both in vitro and in vivo.

To map the SlSPX2‐interacting domain of SlPHR3, Y2H assays using full‐length and truncated SlPHR3 fused to the AD domain, together with full‐length SlSPX2 fused to the BD domain, were performed. Results indicated that the NTD of SlPHR3 mediates binding to SlSPX2 (Figure 5F). Deletion of the NTD completely abolished the interaction, whereas removal of the MYB or coiled‐coil (CC) domain had no significant effect (Figure 5F), demonstrating that the NTD is both necessary and sufficient for the SlPHR3–SlSPX2 interaction.

Notably, the NTD of SlPHR3 is also required for its interaction with SlMED25, as demonstrated previously (Figure 3E). This overlapping domain dependency suggests that SlPHR3, SlSPX2, and SlMED25 operate within an integrated regulatory module to fine‐tune PSR in tomato.

SlSPX2 acts as a corepressor of SlPHR3 to negatively modulate PSR in tomato

To elucidate the molecular and physiological functions of SlSPX2 in tomato PSR, slspx2 knockout mutants using the CRISPR/Cas9 genome editing system were generated. Molecular characterization via sequence alignment identified a 1‐bp insertion in slspx2‐32 and a 1‐bp deletion in slspx2‐43 within the SlSPX2 ORF; both mutations triggered frameshifts and introduced premature stop codons, resulting in loss‐of‐function alleles (Figure S8). SlSPX2 overexpression lines (35S::SlSPX2‐GFP) were constructed and designated as SlSPX2‐OE‐3 and SlSPX2‐OE‐4. RT‐qPCR validation confirmed that SlSPX2 transcript levels were substantially upregulated in these lines relative to WT (Figure S9). Phenotypic analysis under –Pi showed that SlSPX2‐OE plants suffered more severe inhibition of shoot height, fresh weight, and Pi content than WT, whereas slspx2 mutants exhibited reduced growth retardation and more robust Pi homeostasis compared with WT (Figure 6A–D), demonstrating that SlSPX2 negatively regulates PSR‐associated growth and Pi status.

Figure 6.

SlSPX2 acts as a corepressor of SlPHR3 to negatively regulate the tomato PSR

(A) Seedlings of WT, slspx2, and SlSPX2‐OE grown under +Pi or −Pi. Scale bars = 10 cm. (B–D) Inhibition rates of shoot height (B), fresh weight (C), and Pi content (D) in WT, slspx2, and SlSPX2‐OE seedlings under +Pi and −Pi. Data are means ± SD (n ≥ 5 plants). (E) Schematic diagram showing the constructs used in the transient expression assays in (F). (F) Transient expression assays in Nicotiana benthamiana leaves showing that activation of the SlPAP26 by SlPHR3 was reduced by SlSPX2. The ProSlPAP26::LUC reporter was coinfiltrated into N. benthamiana leaves along with the indicated effector constructs. Data are means ± SD (n = 8 leaves). Asterisks indicate significant differences (*P < 0.05, **P < 0.01, ****P < 0.0001, Student's t‐test) (B–D). (F) Different lowercase letters indicate significant differences, as determined by two‐way ANOVA with Tukey's test, P ＜ 0.05.

We next tested whether SlSPX2 represses SlPHR3‐mediated transcriptional activation. Transient expression of SlPHR3 with the proSlPAP26::LUC reporter in N. benthamiana leaves induced a marked increase in luciferase activity, but this activation was significantly impaired by co‐expression of SlSPX2 (Figure 6E, F). No change in reporter activity was detected upon expression of SlSPX2 alone. Collectively, these data establish that SlSPX2 acts as a corepressor of SlPHR3 to fine‐tune tomato PSR.

SlSPX2 and SlMED25 engage in competitive binding to SlPHR3

Given that both SlSPX2 and SlMED25 interact with SlPHR3 via its NTD domain (3, 5), and both Y2H and LCI assays demonstrate no physical interaction between SlSPX2 and SlMED25 (Figure S10), it was postulated that their binding to SlPHR3 is mutually exclusive. To test this, we first performed yeast three‐hybrid (Y3H) assays. The full‐length CDSs of SlMED25 and SlSPX2 were separately cloned into multiple cloning sites I (MCS I) and II (MCS II) of the pBridge vector, generating the pBridge‐SlMED25‐SlSPX2 construct. This vector was co‐transformed with the SlPHR3‐AD fusion construct into yeast. On SD/–Leu/–Trp/–His selective medium, robust interaction between SlPHR3 and SlMED25 was evident; however, induction of SlSPX2 expression by shifting to SD/–Leu/–Trp/–His/–Met medium markedly attenuated the SlPHR3‐SlMED25 interaction (Figure 7A). We further constructed the reciprocal pBridge‐SlSPX2‐SlMED25 vector and conducted parallel Y3H assays with SlPHR3‐AD. The detectable SlSPX2‐SlPHR3 interaction observed on SD/–Leu/–Trp/–His medium was significantly impaired upon induction of SlMED25 expression under SD/–Leu/–Trp/–His/–Met conditions (Figure 7A). These findings demonstrate that SlMED25 and SlSPX2 mutually interfere with one another's binding to SlPHR3; the differential inhibitory effects additionally suggest that SlSPX2 may have a higher binding affinity for SlPHR3 than SlMED25.

Figure 7.

SlSPX2 and SlMED25 compete for binding to SlPHR3

(A) Y3H assays showing that the addition of SlSPX2 dramatically reduced the SlPHR3‐SlMED25 interaction (Top two panels) and the addition of SlMED25 reduced the SlPHR3‐SlSPX2 interaction (Bottom two panels). Yeast cells cotransformed with pGADT7‐SlPHR3 and pBridge‐SlMED25‐SlSPX2 were dropped onto SD/−Trp/−Leu (SD/−2) and SD/−Trp/−Leu/−His (SD/−3) media to assess the SlPHR3‐SlMED25 interaction. The cotransformed yeast cells were dropped onto SD/−Trp/−Leu/−His/−Met (SD/−4) medium to induce expression of SlSPX2. (Bottom two panels) Yeast cells cotransformed with pGADT7‐SlPHR3 and pBridge‐SlSPX2‐SlMED25 were dropped onto SD/−2 and SD/−3 media to assess the SlPHR3‐SlSPX2 interaction. The cotransformed yeast cells were dropped onto SD/−4 medium to induce SlMED25 expression. (B, C) In vitro quantitative pull‐down assays showing that SlSPX2 and SlMED25 compete for binding to SlPHR3. Constant amounts of SlPHR3‐MBP with either SlMED25‐GST (B) or SlSPX2‐His (C) were used. The competing protein, SlSPX2‐His (B) or SlMED25‐GST (C), was titrated in increasing amounts as indicated. SlMED25‐GST and SlSPX2‐His were pulled down by SlPHR3‐MBP immobilized on an anti‐MBP resin. Proteins were eluted and analyzed using the indicated antibodies. The numbers below the bands represent the band intensity values quantified using ImageJ software. (D) A proposed working model showing the role of the SlSPX2‐SlPHR3‐SlMED25 module in tomato PSR. Experiments were repeated at least three times independently with similar results in (A–C).

To further validate this competitive binding model, we performed in vitro pull‐down assays using purified recombinant proteins (SlPHR3‐MBP, SlMED25‐GST, and SlSPX2‐His). When increasing concentrations of SlSPX2‐His were added to a preformed SlPHR3‐MBP/SlMED25‐GST complex, the SlSPX2‐SlPHR3 interaction was dose‐dependently enhanced, whereas the SlMED25‐SlPHR3 interaction was progressively diminished (Figure 7B). In the reciprocal experiment, gradient concentrations of SlMED25‐GST were added to the SlSPX2‐His/SlPHR3‐MBP mixture; notably, no detectable interference with the SlSPX2‐SlPHR3 interaction was observed at low SlMED25‐GST concentrations (Figure 7C). These in vitro data confirm that the SlSPX2‐SlPHR3 interaction exhibits stronger binding affinity than the SlMED25‐SlPHR3 interaction.

DISCUSSION

Recent years have witnessed significant advances in research on PSR in tomato. MYB‐CC transcription factors, including SlPHR1, SlPHR3 (SlPHL1), SlPHR4, SlPHR10, SlPHR11, and SlPHR12, have been shown to be involved in PSR in tomato (Liao et al., 2022; Wu et al., 2023; Lin et al., 2025). Although functional redundancy exists among MYB‐CC transcription factors, AtPHR1 in Arabidopsis and OsPHR2 in rice are both core transcription factors in PSR of their respective species. Currently, it remains unclear which of the aforementioned tomato PHR family members serves as the core transcription factor regulating tomato PSR. Our study revealed that SlPHR3 shares the highest homology with AtPHR1 (Figure S1). Under –Pi stress, the slphr3 single mutant exhibited significantly greater reductions in plant height, phosphate content, and fresh weight compared to WT. Meanwhile, the expression levels of classic PSR marker genes induced by –Pi, such as SlSPX1, SlSPX2, SlPAP26, and SlPT1, were significantly lower in the slphr3 mutant than in WT. Notably, SlPHR3 is able to directly activate the expression of SlSPX1 and SlPAP26 (Figure 1). These phenotypic and molecular traits are highly congruent with the functional characteristics of AtPHR1 and OsPHR2 (Rubio et al., 2001; Zhou et al., 2008), thus indicating that SlPHR3 functions as the key transcription factor regulating PSR in tomato.

In Arabidopsis, PHR1 interacts with MYC2 to coordinately regulate –Pi‐induced jasmonic acid (JA) signaling (He et al., 2023). Our previous work has demonstrated that MED25 interacts with MYC2, the core transcription factor of JA signaling. MED25 recruits Pol II, epigenetic components, and other transcription cofactors to promoters of MYC2 target genes to precisely regulate the transcription initiation of JA signaling in Arabidopsis. And most importantly, one‐third of PHR1 targets are also directly bound by MYC2 (Data not published). Based on these, we speculate that PHR1 may share the same transcriptional machinery components with MYC2. Therefore, we examined the interaction between SlMED25 and SlPHR3 and found that they can directly interact (Figure 3). Genetic data showed that SlMED25 is a positive regulator of PSR in tomato (Figure 2). Further studies revealed that SlMED25 is enriched at the promoter regions of PSR genes, and the regulation of PSR gene expression by SlMED25 is dependent on SlPHR3 (Figure 4B). All these demonstrate that SlMED25 acts as a transcriptional coactivator of SlPHR3. Besides SlMED25, previous studies have shown that MED16 is also involved in PSR in Arabidopsis (Raya‐Gonzalez et al., 2021). Pi starvation inhibits primary root elongation; notably, the root length of med16 mutants exhibits reduced sensitivity to –Pi, accompanied by an attenuated local PSR in root tips. In contrast, the slmed25 mutant shows a significantly higher fresh weight inhibition rate under Pi starvation, with markedly downregulated expression levels of PSR‐related genes (Figure 2A–H). Furthermore, MED16 directly interacts with MED25 and enhances MED25 protein stability, thereby activating JA signaling (Wu et al., 2025). This indicates that MED16 and MED25 synergistically regulate the expression of PSR‐related genes, while exerting opposite functions in mediating the growth inhibition caused by –Pi. Within the Mediator complex, MED25 directly interacts with MED16. Combined with previous studies and our present results, both proteins positively regulate PSR, yet they exert opposite regulatory functions in the growth and development inhibition triggered by –Pi. The underlying molecular mechanism may be associated with the dynamic composition of Mediator complex subunits between shoots and root tips, or it may result from differences in tissue‐specific transcriptional regulation or protein stability control, which warrants further in‐depth investigation. However, it is important to note that the regulatory roles of MED16 and MED25 in PSR‐mediated plant growth and developmental changes may exhibit interspecific differences. Accordingly, future studies should systematically dissect their biological functions in the same plant species to clarify the conservation and specificity of their regulatory mechanisms.

When facing nutrient deficiency or environmental stress, plants initiate transcriptional reprogramming to redirect resources from growth toward stress responses. However, once the stress is alleviated, plants need to promptly terminate stress responses and resume growth (He et al., 2022; Hou and Xu, 2025). As PHR proteins are the core transcription factors involved in PSR in plants, the precise activation and inhibition of their transcriptional activity are crucial for plants to accurately regulate PSR; however, the underlying regulatory mechanism remains largely unclear. We found that the transcriptional activator SlMED25 competes with the transcriptional repressor SlSPX2 for binding to SlPHR3. This competitive interaction enables plants to flexibly regulate the initiation and repression of PSR gene transcription in accordance with dynamic changes in intracellular Pi levels, thereby efficiently coordinating the balance between stress adaptation and growth and development.

In summary, these results definitively demonstrate that SlSPX2 and SlMED25 compete for binding to SlPHR3, with SlSPX2 displaying a superior binding affinity for SlPHR3 relative to SlMED25. This competitive binding module constitutes a central molecular switch that underpins the dynamic regulation of PSR in tomato, with distinct functional outcomes under differing Pi statuses (Figure 7D): (1) Pi‐replete status: High Pi levels stabilize the SlSPX2‐SlPHR3 complex; the robust SlSPX2‐SlPHR3 interaction blocks SlPHR3 from associating with SlMED25 and recruiting the Pol II transcription machinery, consequently suppressing PSR to ensure that plants do not mount unnecessary stress responses under Pi‐sufficient conditions. (2) Pi‐starved status: Low Pi levels disrupt the SlSPX2‐SlPHR3 complex, releasing functional SlPHR3; the freed SlPHR3 then forms a SlPHR3‐SlMED25‐Pol II complex to drive the transcription of PSR target genes, thereby activating adaptive pathways that enhance Pi acquisition and utilization. This regulatory mechanism allows plants to precisely coordinate the binding status of SlPHR3‐interacting proteins, achieving a homeostatic balance between basal growth demands and Pi starvation tolerance.

MATERIALS AND METHODS

Plant materials and growth conditions

The tomato cultivar Solanum lycopersicum Ailsa Craig (AC; LA2838A), provided by the Tomato Genetics Resource Center (TGRC; https://tgrc.ucdavis.edu), was used as the wild type (WT) and genetic background for all mutants and transgenic lines in this study. The SlMED25‐AS and SlMED25‐GFP genotypes have been previously described (Liu et al., 2019). The slphr3 and slspx2 mutants, as well as the SlPHR3‐GFP and SlSPX2‐GFP overexpression lines, were generated in this study. The SlMED25‐GFP transgenic plants were introduced into the slphr3 mutant background via genetic crossing, and homozygous plants were confirmed by genotyping. Tomato seeds were surface‐sterilized and germinated by incubation in sterile water at 25°C for 48 h.

Seedlings were then grown on half‐strength Murashige and Skoog medium, supplemented with 1% (w/v) sucrose and solidified with 0.8% (w/v) agar (pH 5.8), under continuous white light (200 μmol photons·m⁻²·s⁻¹) at 26°C for 5 d. Then the seedlings were transplanted into pots filled with pure vermiculite and irrigated with the respective Hoagland solutions with different Pi levels. The plants were maintained under controlled environmental conditions with a 16‐h light (26°C ± 2°C)/8‐h dark (20°C ± 2°C) photoperiod. For low Pi treatment, a Hoagland nutrient solution system (pH 5.8) was employed, with Pi (KH₂PO₄) concentrations set at 0.05 mM (low Pi, –Pi) and 1.0 mM (normal Pi, +Pi). The nutrient solutions were added every 3 d. After 21 d of treatment, plant phenotypes were observed, and multiple physiological indices were measured.

Plasmid construction and plant transformation

To generate the SlPHR3‐GFP and SlSPX2‐GFP constructs, the full‐length coding sequence (CDS) of SlPHR3 and SlSPX2 was amplified by PCR and cloned into the pK7FWG2.0 vector downstream of the Cauliflower mosaic virus (CaMV) 35S promoter to generate SlPHR3–GFP and SlSPX2–GFP fusion constructs. The primers utilized in this cloning process are detailed in Table S1. The resulting plasmids, designated pro35S::SlPHR3‐GFP and pro35S::SlSPX2‐GFP, were then transferred into AC via Agrobacterium‐mediated transformation (Du et al., 2014). Transgenic plants were screened on kanamycin‐containing medium, and homozygous T2 lines were subsequently employed for phenotypic and molecular analyses.

CRISPR/Cas9‐mediated mutation

The mutants of slphr3‐cr and slspx2‐cr were constructed via CRISPR/Cas9 technology (Deng et al., 2023). To generate guide RNAs (gRNAs) for CRISPR/Cas9‐mediated editing of SlPHR3 and SlSPX2, two 19‐bp sequences within the coding region of each gene were selected. These gRNA sequences were PCR‐amplified using the pHSE401 vector as a template, with forward and reverse primers incorporating the respective 19‐bp targeting sequences (Table S1). The tomato U6‐26‐SlPHR3‐gRNA expression cassette and the CRISPR/Cas9 binary vector pCBC‐DT1T2_tomatoU6 were both digested with the BsaI restriction enzyme. The digested cassette was then ligated into the binary vector, resulting in the final constructs pCBC‐DT1T2_tomatoU6‐SlPHR3 and pCBC‐DT1T2_tomatoU6‐SlSPX2. These constructs were introduced into AC through Agrobacterium tumefaciens‐mediated transformation (Du et al., 2014). Mutations induced by CRISPR/Cas9 were identified by PCR amplification and subsequent DNA sequencing of the target regions. All primers employed for plasmid assembly are provided in Table S1. In the T2 generation, plants that carried mutations in SlPHR3 or SlSPX2 but lacked the Cas9 transgene were selected for subsequent experiments.

Growth and physiological analysis of tomato plants

The inorganic Pi content in plants was measured using a modified version (Chiou et al., 2006). Seedlings were cultivated for 21 d under different Pi conditions (+Pi and –Pi). Phenotypes were evaluated by measuring shoot fresh weight, plant height, and Pi content of leaves. For Pi quantification, frozen leaf samples were homogenized in 1 mL of 10% (w/v) perchloric acid, followed by a 10‐fold dilution with 5% (w/v) perchloric acid. After centrifugation at 10,000 × g for 10 min, the supernatant was collected for Pi assay via the molybdenum blue method. Briefly, 1 mL of each sample was mixed with 2 mL of a working solution composed of six parts solution A (0.4% ammonium molybdate in 0.5 M H₂SO₄) and one part solution B (10% ascorbic acid). The mixture was incubated at 42°C for 20 min, cooled on ice, and absorbance was measured at 820 nm using a spectrophotometer. Pi concentrations were determined based on the absorbance values according to a previously described protocol (Dong et al., 2019). Inhibition rate of plant height, fresh weight, and Pi content was calculated by the following equation:

Inhibition rate of shoot height

$$=\frac{\text{The shoot height }\left(+\text{Pi condition}\right)-\text{The shoot height}\left(-\text{Pi condition}\right)}{\text{The shoot height }\left(+\text{Pi condition}\right)}\times 100\%$$

Inhibition rate of fresh weight

$$=\frac{\text{The fresh weight }\left(+\text{Pi condition}\right)-\text{The fresh weight }\left(-\text{Pi condition}\right)}{\text{The fresh weight }\left(+\text{Pi condition}\right)}\times 100\%$$

Inhibition rate of Pi content

$$=\frac{\text{The Pi content }\left(+\text{Pi condition}\right)-\text{The Pi content }\left(-\text{Pi condition}\right)}{\text{The Pi content }\left(+\text{Pi condition}\right)}\times 100\%$$

For the analysis of growth phenotypes, eight biological replicates were employed, with one plant designated as a single replicate. For the determination of Pi contents, five biological replicates were used, following the same replicate design (one plant per replicate).

RNA extraction and gene expression analysis

For RT‐qPCR analysis, total RNA was extracted from leaves or other relevant tissues. Seedlings were first cultured on 1/2MS medium (Pi‐sufficient, +Pi) for 3 d, then transferred to 1/2MS medium supplemented with +Pi or deprived of Pi (–Pi) and incubated for 48 h prior to tissue sampling. Total RNA was extracted using a TransZol Up Plus RNA Kit (TransGen) according to the manufacturer's instructions. For RT‐qPCR analysis, cDNA was synthesized from 1 μg of total RNA using TransScript® One‐Step gDNA Removal and cDNA Synthesis SuperMix (TransGen). Amplification was performed on a Roche 480 instrument using the TransStart Tip Green qPCR SuperMix kit (TransGen), and the relative expression levels of target genes were calculated using ACTIN2 as the internal reference (Du et al., 2017). Three technical replicates were included for each biological sample, and the results are presented as the mean of three independent biological replicates, with error bars indicating the standard deviation. Each replicate represented a pool of leaves or other tissues collected from ten seedlings grown under +Pi or –Pi. All primers used in this study are provided in Table S1.

Y2H assays

Yeast two‐hybrid (Y2H) analyses were conducted with the Matchmaker Gold Yeast Two‐Hybrid System (Clontech). The CDSs of SlMED25 and SlSPX2 were amplified from cDNA and inserted into the pGBKT7 vector, while the full‐length SlPHR3 CDS was cloned into pGADT7 to examine its interaction with these proteins. For domain mapping of SlMED25, its full‐length and truncated versions were subcloned into pGBKT7, with SlPHR3 in pGADT7. Similarly, to identify SlPHR3 regions mediating interactions with SlMED25 or SlSPX2, full‐length and derivative sequences of SlPHR3 were ligated into pGADT7, while SlMED25 or SlSPX2 CDS was placed in pGBKT7. All primers used are provided in Table S1. Constructs containing SlMED25 or SlSPX2 together with SlPHR3 were co‐introduced into the Saccharomyces cerevisiae Y2HGold strain. Empty vectors served as negative controls. A 5 μL aliquot of each transformed yeast suspension (OD₆₀₀ = 1.0) was spotted onto: SD/–Leu/–Trp medium to verify the transformation; SD/–His/–Leu/–Trp and SD/–Ade/–His/–Leu/–Trp media to assess the protein–protein interactions. Plates were incubated at 30°C for 72 h, and all assays were performed in three independent replicates.

Y3H assays

The full‐length CDS of SlPHR3 was inserted into the pGADT7 vector to generate an AD fusion construct. For the yeast three‐hybrid (Y3H) analysis, the pBridge vector (Clontech) was utilized to generate dual‐expression constructs. In the pBridge‐SlMED25‐SlSPX2 construct, the SlMED25 CDS was ligated into MCS I, generating a fusion with the GAL4 DNA‐binding domain (BD), while the SlSPX2 CDS was introduced into MCS II to enable methionine‐repressible expression. Similarly, the pBridge‐SlSPX2‐SlMED25 construct was generated by cloning SlSPX2 into MCS I and SlMED25 into MCS II. Yeast three‐hybrid assays were performed using the MATCHMAKER GAL4 Two‐Hybrid System (Clontech). All plasmid combinations were co‐transformed into the Saccharomyces cerevisiae strain AH109. Transformants were initially selected on SD/–Leu/–Trp (SD/–2) medium. To evaluate the interaction between SlPHR3 and SlMED25 in the absence of the third protein, yeast cells were plated on SD/–Leu/–Trp/–His (SD/–3) medium. For assays requiring the expression of the bridge protein (SlSPX2 or SlMED25), cells were cultured on SD/–Leu/–Trp/–His/–Met (SD/–4) medium. All plates were incubated at 30°C for 3 d before interaction outcomes were assessed.

LCI assays

Split‐luciferase complementation imaging (LCI) assays were performed in N. benthamiana leaves following an established protocol. The CDS of SlMED25 and SlSPX2 were inserted into the pCAMBIA1300‐cLUC vector, whereas the SlPHR3 CDS was subcloned into the pCAMBIA1300‐nLUC vector (Table S1 for primers). Each resulting plasmid was introduced into Agrobacterium tumefaciens strain GV3101. Transformed agrobacteria were grown in Yeast Extract Peptone (YEP) medium at 28°C for 24 h, then diluted 1:100 (v/v) into fresh YEP medium supplemented with 10 mM MES and 40 μM acetosyringone, and cultured for an additional 16 h. Cells were harvested by centrifugation at 4,000 × g for 10 min and resuspended in 10 mM MgCl₂ containing 0.2 mM acetosyringone to a final OD₆₀₀ of 1.5. After incubating the suspensions at room temperature for 3 h, equal volumes of agrobacteria carrying nLUC and cLUC constructs were mixed and co‐infiltrated into the abaxial side of N. benthamiana leaves. SlSCaBP8‐nLUC and cLUC‐SlULP1 served as negative controls. Infiltrated plants were maintained at 23°C under a 16‐h‐light/8‐h‐dark cycle for 72 h. Prior to imaging, leaves were sprayed with 0.5 mM luciferin and kept in darkness for 3 min. Luciferase activity was visualized using a low‐light cooled CCD camera (NightOWL II LB983 with Indigo software).

In vitro pull‐down assays

To produce MBP‐ or GST‐tagged recombinant proteins, the CDS of SlMED25, SlPHR3, and SlSPX2 were cloned into pMAL‐c2X or pGEX‐4T‐3 vectors, respectively. SlSPX2 was also inserted into pET‐28a for His‐fused protein expression. All primers are listed in Table S1. The resulting plasmids were introduced into Escherichia coli BL21 (DE3) cells. Protein expression was induced with 0.5 mM IPTG, and the MBP‐, GST‐, and His‐tagged fusion proteins were subsequently purified using amylose resin (New England Biolabs), GST bind resin (Thermo Fisher), and Ni‐NTA resin (EedChemExpress), respectively. Band intensity quantification was performed using ImageJ software.

Co‐IP assays

Co‐immunoprecipitation (Co‐IP) assays were conducted in N. benthamiana leaves following a modified protocol from a previous study (Zhang et al., 2018). Total protein was extracted from 500 mg of leaf tissue collected 48 h post‐infiltration by grinding in liquid nitrogen followed by homogenization in 2 mL of ice‐cold IP buffer (50 mM Tris‐HCl (pH 7.5), 150 mM NaCl, 0.1% Triton X‐100, 0.2% NP‐40, 0.6 mM PMSF, and 20 μΜ MG132 with Roche protease inhibitor cocktail). The homogenate was centrifuged at 12,000 × g for 20 min at 4°C, and the supernatant was collected as the total protein extract. For each immunoprecipitation reaction, 500 μg of total protein was incubated with 20 μL of anti‐GFP magnetic beads for 2 h at 4°C with gentle rotation. Control samples included SlMED25‐myc co‐expressed with free GFP, and SlSPX2‐Flag co‐expressed with free GFP. After incubation, the beads were washed five times with 1 mL of IP buffer, and bound proteins were eluted by boiling in 2× SDS loading buffer for 10 min. Both input extracts and immunoprecipitated samples were subjected to immunoblot analysis using anti‐GFP (1:5,000), anti‐MYC (1:3,000), and anti‐Flag (1:3,000) antibodies (TransGen).

ChIP assay

Chromatin immunoprecipitation (ChIP) assays were performed using 6 d‐old seedlings of SlPHR3‐GFP, SlMED25‐GFP, and SlMED25‐GFP/slphr3 grown under +Pi and –Pi biological conditions. For each sample, approximately 2 g of whole seedlings were harvested and subjected to in vivo cross‐linking with 1% (v/v) formaldehyde under vacuum infiltration for 10 min at room temperature. The cross‐linking reaction was quenched by adding glycine to a final concentration of 0.125 M, followed by an additional 5‐min vacuum treatment. The fixed tissues were cryogenically pulverized in liquid nitrogen, and chromatin complexes were extracted in nuclei lysis buffer (50 mM HEPES [pH 7.5], 150 mM NaCl, 1 mM EDTA, 1% SDS, 1% Triton X‐100, 0.1% sodium deoxycholate, 1 mM PMSF, and protease inhibitor cocktail). Chromatin was sheared by sonication to reduce the average DNA fragment size to approximately 500 bp. An input control (50 μL) was removed prior to immunoprecipitation. The remaining chromatin lysate was incubated overnight at 4°C with anti‐GFP antibody (AB290, Abcam), followed by capture with protein G beads (Invitrogen). The beads were sequentially washed with low‐salt buffer (150 mM NaCl, 0.2% SDS, 0.5% Triton X‐100, 2 mM EDTA, 20 mM Tris‐HCl [pH 8.0]), high‐salt buffer (500 mM NaCl, 0.2% SDS, 0.5% Triton X‐100, 2 mM EDTA, 20 mM Tris‐HCl [pH 8.0]), LiCl wash buffer (0.25 M LiCl, 0.5% NP‐40, 0.5% sodium deoxycholate, 1 mM EDTA, 10 mM Tris‐HCl [pH 8.0]), and TE buffer. Immunoprecipitated complexes were eluted using elution buffer (1% SDS, 100 mM NaHCO₃), and cross‐links were reversed by overnight incubation at 65°C. DNA was purified using the QIAquick PCR Purification Kit (Qiagen) and subjected to qPCR analysis. The qPCR employed specific primers targeting regions either containing or lacking the P1BS element within the promoters of SlSPX1 and SlPAP26 (Table S1). ACTIN2 served as a negative control. Enrichment values were calculated by normalizing to the input sample and expressed as fold enrichment.

EMSA

The full‐length CDS of SlPHR3 was amplified via PCR and inserted into the pGEX‐4T‐3 vector to generate a GST‐tagged fusion construct. Recombinant GST‐SlPHR3 protein was expressed in Escherichia coli strain BL21 (DE3) and affinity‐purified to homogeneity using GST‐binding resin. Biotin‐labeled oligonucleotide probes were synthesized with 5′‐end modifications (Invitrogen). Electrophoretic mobility shift assays (EMSAs) were conducted as previously described (Chen et al., 2011). In brief, the biotinylated probes were incubated with purified GST‐SlPHR3 protein for 20 min at room temperature. Protein–DNA complexes were resolved from unbound probes by non‐denaturing polyacrylamide gel electrophoresis (PAGE). As negative controls, mutated versions of the probes were used in which the native P1BS motifs (5′‐GGATATTC‐3′ in SlSPX1 and 5′‐GAATATTC‐3′ in SlPAP26) were replaced with a scrambled sequence (5′‐AAAAAAAA‐3′). All probe sequences are provided in Table S1.

Transient expression assays

Transient transcriptional activity assays were performed using a dual‐luciferase reporter system in N. benthamiana leaves. A 2,177‐bp fragment of the SlPAP26 promoter was isolated from tomato genomic DNA and inserted into the pGreenII 0800‐LUC vector to serve as the firefly luciferase (LUC) reporter. The Renilla luciferase (REN) gene, driven by the CaMV 35S promoter in the same vector, was used as an internal control. Effector constructs consisted of SlPHR3‐GFP, SlMED25‐GFP, and SlSPX2‐GFP under the control of the 35S promoter. All primer sequences are provided in Table S1. The reporter and effector constructs were introduced into Agrobacterium tumefaciens strain GV3101. Bacterial cultures were grown to mid‐log phase, harvested by centrifugation, and resuspended in infiltration medium (10 mM MES, 10 mM MgCl₂, and 0.2 mM acetosyringone, pH 5.6) to an OD₆₀₀ of 0.5. Agrobacterial suspensions containing the reporter and respective effector constructs were mixed in equal volumes and co‐infiltrated into the abaxial side of N. benthamiana leaves. After 48–60 h, LUC and REN activities were quantified using the Dual‐Luciferase® Reporter Assay System (Promega) according to the manufacturer's protocol. The relative transcriptional activity was expressed as the ratio of LUC to REN luminescence. Three independent biological replicates were performed for each construct combination.

Phylogenetic tree construction

Phylogenetic analysis was performed using the MEGA X program with the maximum‐likelihood method. The robustness of the tree topology was assessed with 1,000 bootstrap replicates. The amino acid sequences utilized for the construction of the PHR and SPX phylogenetic trees are provided in [Link], [Link], respectively.

Subcellular localization assay

The CDSs of SlSPX1, SlSPX2, SlSPX3, SlSPX4, and SlSPX5 were amplified and cloned into the GFP‐fusion expression vector pK7FWG2.0. Both the recombinant vector and the empty vector, which carries the 35S::GFP, were introduced into A. tumefaciens strain GV3101 for further infiltration into N. benthamiana leaves. The confocal microscopy LSM 880 was applied to detect the fluorescence signals.

Statistical analysis

All statistical analyses were conducted in GraphPad Prism 8.0. The averages and standard deviations (SD) of all results were calculated. The significance of the difference was examined by a two‐tailed Student's t‐test or a two‐way ANOVA with Tukey's test.

Accession numbers

Sequence data from this article can be found in the Sol Genomics Network Initiative under the following accession numbers: Solyc12g070100 (SlMED25), Solyc05g055940 (SlPHR1), Solyc04g050070 (SlPHR2), Solyc09g072830 (SlPHR3), Solyc06g008200 (SlPHR4), Solyc08g060920 (SlSPX1), Solyc12g009480 (SlSPX2), Solyc01g090890 (SlSPX3), Solyc02g067160 (SlSPX4), Solyc02g088210 (SlSPX5), Solyc09g090070 (SlPT1), Solyc07g007670 (SlPAP26), and Solyc11g005330 (SlACTIN2).

CONFLICTS OF INTEREST

The authors declare no conflicts of interest.

AUTHOR CONTRIBUTIONS

Q.C., C.L. conception and design of the experiments. M.Z., H.H., Y.Z., Y.Z., H.W., and H.G. performed the experiments. X.M., C.S., Q.C., C.L., data analysis. M.Z., L.D., Q.C., and C.L. prepared the manuscript. All authors critically reviewed the study results and unanimously approved the final version of the manuscript.

Supporting information

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.70278/suppinfo

Figure S1. Phylogenetic tree of PHR proteins from Solanum lycopersicum and Arabidopsis thaliana

Figure S2. Generation of slphr3 mutants using CRISPR/Cas9 technology

Figure S3. Generation of SlPHR3 overexpression plants (SlPHR3‐OE)

Figure S4. SlMED25‐mediated PSR gene expression is dependent on SlPHR3

Figure S5. Expression analysis of SlMED25 in different tissues

Figure S6. Phylogenetic analysis of SPX proteins of Solanum lycopersicum and Arabidopsis thaliana

Figure S7. Subcellular localization of SlSPX4‐GFP and SlSPX5‐GFP in Nicotiana benthamiana leaves

Figure S8. Generation of slspx2 mutants via CRISPR/Cas9 technology

Figure S9. Generation of SlSPX2 overexpression plants (SlSPX2‐OE)

Figure S10. No interaction was detected between SlSPX2 and SlMED25

Table S1. List of primers and probes used in this study

Table S2. Amino acid sequences of PHRs in Arabidopsis and tomato

Table S3. Amino acid sequences of SPXs in Arabidopsis and tomato

Biographies

Zhai, M. , Han, H. , Zhang, Y. , Zhao, Y. , Guo, H. , Wang, H. , Sun, C. , Meng, X. , Deng, L. , Chen, Q. , et al. (2026). SlMED25‐SlPHR3‐SlSPX2 module fine‐tunes SlPHR3‐mediated transcriptional activation of phosphate starvation response in tomato. J. Integr. Plant Biol. 68: 3593–3609.