Gibberellin and abscisic acid transporters facilitate endodermal suberin formation in Arabidopsis

doi:10.21203/rs.3.rs-1670556/v1

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Gibberellin and abscisic acid transporters facilitate endodermal suberin formation in Arabidopsis

https://doi.org/10.21203/rs.3.rs-1670556/v1

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published 06 Apr, 2023

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The plant hormone gibberellin (GA) regulates multiple developmental processes. It accumulates in the root elongating endodermis, but how it moves into this cell file and the significance of this accumulation are unclear. Here, we identified a monophyletic clade of NPF transporters required for GA and abscisic acid (ABA) translocation. We demonstrate that NPF2.14 is a subcellular GA/ABA transporter, the first to be identified in plants, facilitating GA and ABA accumulation in the root endodermis to regulate suberization. Further, NPF2.12 and NPF2.13, closely related proteins, are plasma membrane-localized GA and ABA importers that facilitate shoot-to-root GA₁₂ translocation, regulating endodermal hormone accumulation. This work reveal that GA promotes root suberization and that GA and ABA can act non-antagonistically. We demonstrated how a clade of transporters mediates hormone flow while utilizing defined cell-file-specific vacuolar storage at the phloem unloading zone, allowing a hormone slow-release mechanism required for suberin formation in the maturation zone.

The phytohormone gibberellin (GA) is essential for many developmental processes in plants. Among them are seed germination, organ elongation and expansion through cell growth and division, trichome development, the transition from vegetative to reproductive growth, and flower, seed, and fruit development ¹. GAs are produced mainly in the vasculature and move long distances in both acro- and basipetal directions ^2–5. The first reports of GA mobility through the phloem sap appeared over 50 years ago ^6–8. Grafting methods showed that biosynthesis of GA in the shoot can rescue GA biosynthesis mutations in the hypocotyl and root and vice versa ^6–8. In tobacco plants, defoliation results in an internode elongation, cambial activity, and fiber differentiation phenotypes that are similar to treatment with paclobutrazol, a GA biosynthesis inhibitor ⁵. Another example of an organ dependent on an external source of GA are the petals, which require the anthers as their GA source ^9,10. The biosynthesis of active GA is a complex, multi-step process with diverse intermediates ¹¹. Regnault et al. conducted a series of grafting experiments using Arabidopsis thaliana mutant plants compromised at different stages of GA biosynthesis and identified GA₁₂ as the major GA form transported over a long distance through the vasculature ¹². GA₁₂ moves through the xylem in a root-to-shoot manner and in the phloem in a shoot-to-root direction to regulate plant growth ¹². The translocation of GA₁₂ from the root to the shoot is enhanced under ambient temperatures to induce shoot growth ¹³. The transporters that regulate GA long-distance transport in the plant remain unknown.

In recent years, a number of Arabidopsis GA plasma membrane transporters have been identified, two from the SWEET family and several others from the NITRATE TRANSPORTER1/PEPTIDE TRANSPORTER (NPF) family ^14–17. There are 53 NPFs in Arabidopsis, divided into eight subfamilies. These proteins are capable of transporting a large variety of substrates such as nitrate, glucosinolates, abscisic acid (ABA), auxin, and GAs ^14,18,19. Like auxin, GA is subjected to the ion-trap mechanism, limiting its ability to move out of cells. The existence of GA efflux transporters is therefore predicted to allow GA cell-to-cell movement ²⁰; however, no GA efflux transporters have been discovered ²¹.

ABA, which regulates growth and stress responses, has been long thought to act antagonistically to GA in processes such as seed germination, seed maturation, and dormancy and in responses to external cues ^22–24. Both GA and ABA induce developmental responses specifically from the endodermis ^25–27. GA accumulates at high levels in the endodermal cells of the root elongation zone ²⁸. The process is dependent on the activity of NPF3.1 (NPF3), a member of the NPF family, which has been shown to act as a dual-specificity GA and ABA importer ^16,29. The endodermis, the innermost cortical layer that surrounds the central vasculature, goes through two phases of differentiation: The first step involves polar localized lignin deposition, which results in the formation of the casparian strips, and the second is the deposition of suberin as a lamella below the primary cell wall ³⁰. The casparian strips restrict apoplastic diffusion of water and nutrients into the vascular tissues, whereas suberin limits backflow of nutrients from the stele ^31,32. The physiological relevance of GA accumulation in the endodermis remains unknown.

With this study, we identified a monophyletic clade of NPF transporters that orchestrate GA₁₂ long-distance shoot-to-root translocation and promote bioactive ABA and GA movement from the vasculature to the endodermis, which is required for endodermal suberization, a phenotype rescued by applying either ABA or GA. The importance of transporter distributions was evaluated via a multicellular mathematical model that revealed an intriguing slow-release mechanism wherein high levels of GA and ABA in the phloem unloading zone are loaded into pericycle vacuoles for continued induction of suberin formation in the maturation zone. Together, our findings reveal the mechanism that facilitates long-distance shoot-to-root movement of GA₁₂ and explain how bioactive GA₄ and ABA are transported from the vasculature to the endodermis to mediate endodermal suberization.

NPF2.14 is a tonoplast-localized GA and ABA transporter

To identify the missing GA exporters in plants, we tested whether NPF proteins were capable of GA₄ export in Xenopus laevis oocyte-based transport assays. Other GA transporters identified to date import GA ^14,33. The transport assays investigate whether NPFs lead to efflux of GA that is loaded into oocytes by three different approaches, namely diffusion, injection and import by a GA importer. Oocytes expressing various NPFs or control oocytes injected with water were exposed to an array of membrane-permeable GAs (GA₄, GA₇, GA₉ and GA₁₂) ³⁴. The screen has identified NPF2.14 as a potential GA exporter. At the end of assay, NPF2.14-expressing oocytes contained significantly reduced GA levels compared to control oocytes (Fig. 1A). This suggests that NPF2.14 facilitates the export activity of GAs out of oocytes. This was confirmed in an injection-based assay in which GA₃, a non-membrane-permeable form of GA, was injected directly into the oocyte. At the end of the assay, there was a lower GA₃ content in NPF2.14-expressing oocytes compared to control oocytes (Fig. 1B). Co-expression of both NPF2.14 RNA and NPF4.1, which encodes a known GA importer, led to a reduced accumulation of non-membrane permeable GA₃ compared to oocytes expressing only NPF4.1 (Fig. 1C), further supporting that NPF2.14 exports GA out of Xenopus laevis oocytes.

It was previously shown that NPF3.1 transports both GA and ABA in Xenopus laevis oocytes ¹⁶. To test whether NPF2.14 also has broad substrate specificity, control and NPF2.14-expressing oocytes were exposed to ABA. NPF2.14-expressing oocytes accumulated less ABA than did control oocytes (Fig. 1D), indicating that NPF2.14 has dual-substrate transport activity. Several NPF transporters, including NPF6.3, also transport nitrate ¹⁸, but NPF2.14 displayed no nitrate transport activity in oocyte assays (Fig. 1E).

Wulff et al. recently showed that NPF7.3 lowers the cytoplasmic pH in Xenopus laevis oocytes, which can indirectly influence the accumulation equilibrium of weak acids such as GA and ABA ³⁴. In order to assess whether NPF2.14 has a similar activity, we measured the intracellular oocyte pH using a proton-selective three-electrode voltage clamp setup. We showed that, unlike NPF7.3, NPF2.14 expression in the oocyte does not alter the internal oocyte pH (Fig. 1F). Another factor that theoretically can lead to a false-positive GA export result is alteration of membrane potential. Therefore, we measured the membrane potential of control and NPF2.14-expressing oocytes using a two-electrode voltage clamp setup. The membrane potential of control and NPF2.14-expressing oocytes were both approximately − 15 mV (Fig. 1G). When oocytes were subjected to GA₄ for 60 min prior to membrane potential measurement, less GA was observed in NPF2.14-expressing oocytes than control oocytes (Fig. 1G). Thus, NPF2.14 does not shift oocyte membrane potential.

Many NPF proteins, including NPF2.14, contain the ExxE[K/R] motif ³⁵, which is involved in coupling substrate transport to the proton gradient across membranes ³⁶. Involvement of the ExxE[K/R] motif in NPF2.14-mediated effluxes would suggest antiporter function. To assess the involvement of the ExxE[K/R] motif, we generated C-terminal YFP-tagged NPF2.14 mutants substituted at each of the three charged residues with a polar but uncharged residue. The YFP-tag alone did not influence the apparent GA₄ transport by the wild-type NPF2.14 (Fig. 1H). When any of the charged residues of the ExxE[K/R] motif were replaced with the polar uncharged Gln residue, no significant difference in GA₄ transport was observed compared to wild-type NPF2.14 (Fig. 1H). Thus, the GA₄ transport mediated by NPF2.14 seems to be ExxE[K/R] motif independent.

In order to test whether the Xenopus laevis oocyte GA transport data is physiologically relevant in planta, we isolated a homozygous T-DNA knockout line for NPF2.14. The single npf2.14 mutant did not show significant shoot or root growth phenotypes (Sup. Figure 1A-C). Several NPF family members transport nitrate, including NPF6.3 ³⁷. Thus, despite having shown that NPF2.14 does not transport nitrate in oocytes (Fig. 1D), we checked whether npf2.14 mutants display an impaired growth on low nitrate media. npf2.14 T-DNA insertion mutants did not have a visible growth phenotype and did not differ from Col-0 plants under low nitrate conditions (Sup. Figure 1D). To determine whether NPF2.14 is involved in GA distribution and accumulation in the root, we tested whether the distribution of a fluorescently tagged GA₃ compound (GA₃-Fl) was affected in the loss-of-function line. GA₃-Fl has been developed in our lab to serve as a stable, bioactive reporter to study GA movement/accumulation in planta ²⁸. Accumulation of GA3-Fl in the endodermis was visible in the Col-0 plants, as previously shown ²⁵. The npf2.14 mutants displayed a significantly stronger signal compared to the Col-0 control (Fig. 1I). This enhanced accumulation was restored to normal levels when expressing NPF2.14 driven by its native promoter (pNPF2.14:NPF2.14-GFP) on the background of the npf2.14 T-DNA line (Fig. 1I), indicating that loss of NPF2.14 affects GA3-Fl distribution in the plant. In agreement with this result, npf2.14 mutants accumulated significantly higher levels of GA₄ in their roots (Fig. 1J).

In order to study the subcellular localization of NPF2.14, we generated and imaged 35S:NPF2.14-YFP lines. Interestingly, NPF2.14 localized to the tonoplast vacuole membrane (Fig. 1K). Therefore, although we had hypothesized that NPF2.14 was a GA exporter that transported GA from inside the cytosol to the apoplast, the protein is instead a tonoplast-localized transporter. To the best of our knowledge, this is the first report of a sub-cellular GA/ABA transporter.

NPF2.14 regulates suberin formation in the root endodermis

To characterize NPF2.14 expression patterns in the plant, we generated NLS-YFP and GUS reporter lines driven by the NPF2.14 promoter. Confocal imaging of the NLS-YFP lines indicated that NPF2.14 is expressed only in the pericycle of the root mature zone, mainly in the phloem poles, and not in the meristematic zone (Fig. 2A, Sup. Fig. 2A). In addition, GUS staining showed expression in the shoot vasculature in seedlings (Sup. Fig. 2B). In the mature stages, the NPF2.14-driven reporter was expressed in the periderm (Sup. Fig. 2C). The pericycle is a deep layer of post-embryonic meristematic cells encircling the vascular tissue ³⁸. In the root, it is required for lateral root emergence ³⁹, xylem loading ⁴⁰ and phloem unloading ⁴¹. Later stages give rise to the periderm, which serves as the outer protective layer when the surrounding tissue is sloughed off ³⁸. Both the pericycle and the cork, which is the outermost cell layer of the periderm are suberinized tissues ⁴².

The expression of the reporter driven by the NPF2.14 promoter in tissues that undergo suberization, taken together with the ability of NPF2.14 to transport ABA, which has been previously shown to regulate root suberization ⁴³, led us to hypothesize that this transporter might facilitate root suberization.

To test this, we analyzed suberization in npf2.14 T-DNA mutants using Nile red and Fluorol yellow, which are suberin dyes ^44,45. Suberization commences in the endodermis of the upper part of the maturation zone of the root, and, as the plant matures more cells undergo suberization ⁴⁶. Quantification of Nile red and Fluorol yellow fluorescence intensity in the uppermost part of 5-day-old roots revealed that the mutant npf2.14 plants had significantly lower levels of endodermal suberization than Col-0 plants (Fig. 2B, Sup. Fig. 3).

Suberin is a complex polyester based on glycerol and long-chain α,ω-diacids and ω-hydroxyacids ⁴⁷, which is primarily found in structures such as the periderm, endodermis, and seed coat ³⁰. To examine changes in root suberin composition between npf2.14 and Col-0, we analyzed their suberin monomer profiles via gas chromatography-mass spectrometry (GC-MS). We found significant reductions in ferulic acid, the predominant aromatic component of suberin, as well as in C22 fatty acid and C18:1(9) ω-hydroxyacid, two of the most abundant suberin building blocks in the Arabidopsis root endodermis (Fig. 2C). In addition, C18 ester was lower in npf2.14 roots. These reductions accompanied by lower levels of other monomers resulted in 35% less total suberin contents in the mutant roots (Fig. 2C). Overall, the findings provide several lines of evidence that alteration of NPF2.14 has a substantial effect on suberin deposition in the root endodermis.

Similar to the npf2.14 mutant, npf3.1 mutant plants, which have been shown to have impaired GA and ABA delivery to the endodermis ¹⁶, displayed reduced suberization levels compared to Col-0 plants (Sup. Fig. 4). GA and ABA treatments completely rescued the npf2.14 mutant suberization levels (Fig. 2B).

Endodermal suberization is regulated by ABA perception, both under normal and stress conditions ^43,48. ABA treatment was previously reported to induce endodermal suberization ⁴⁹. In agreement with this report, ABA significantly upregulated the Nile red fluorescence intensity in the root endodermis and rescued the reduced suberization observed in the aba2-1 mutant, which is deficient in ABA biosynthesis ⁴⁹ (Fig. 2D). To the best of our knowledge, GA has not been previously associated with endodermal suberization. To establish whether or not GA regulates root suberization in Arabidopsis seedlings, we quantified suberization levels in GA-treated wild-type and ga1-13 mutant plants using Nile red and Fluorol yellow dyes. GA1 catalyzes the first committed step in the GA biosynthetic pathway ⁵⁰. GA treatment to wild-type caused significant induction in endodermal root suberization (Fig. 2E, Sup. Fig. 5), and the GA biosynthesis mutant ga1-13 displayed a significant reduction in suberization levels, which was restored by the exogenous application of GA₃ (Fig. 2E). GA and ABA have long been thought to have completely antagonistic functions ⁵¹. Our results indicate that this antagonistic activity is more complex and that GA and ABA enhance root suberization. Taken together, our data suggest that NPF2.14 is a pericycle-specific GA and ABA transporter, localized to the tonoplast, and involved in regulating GA and ABA accumulation in the endodermis to promote suberization.

NPF2.12 and NPF2.13 are plasma membrane-localized GA and ABA importers

NPF2.12 and its close paralog NPF2.13 form a monophyletic phylogenetic clade with NPF2.14 (Fig. 3A, Sup. Fig. 6). We hypothesized that due to this proximity on the phylogenetic tree, NPF2.12 and NPF2.13 might also contribute to GA and ABA accumulation in the endodermis. Both transporters were previously characterized as low-affinity nitrate transporters ^52,53 and more recently, were shown to promote GA import activity in heterologous systems ¹⁴^,³⁴. Neither have been characterized in plants as GA or ABA transporters. To test for direct GA transport activity of NPF2.12 and NPF2.13, we performed Xenopus laevis oocyte-based transport assays. Oocytes expressing NPF2.12 or NPF2.13 accumulated significantly higher levels of GA₁, GA₃, GA₄, GA₇, GA₉, GA₁₂, GA₁₉, and GA₂₄compared to control oocytes over the course of 60 min (Fig. 3B, Sup. Fig. 7A). This suggests that NPF2.12 and NPF2.13 are promiscuous GA transporters. Both NPF2.12- and NPF2.13-expressing oocytes also had higher levels of ABA accumulation than controls (Fig. 3C).

NPF2.12 and NPF2.13 both contain an ExxE[K/R] motif, which couples the substrate transport to the proton gradient ³⁶. To test whether NPF2.12 and NPF2.13 substrate transport activity is coupled with external proton concentration, NPF2.12- and NPF2.13-expressing oocytes were treated with membrane-impermeable GA₁ in solutions ranging from pH 5 to 7 in 0.5 pH unit increments. In both NPF2.12- and NPF2.13-expressing oocytes, GA₁ accumulation increased as pH was lowered. (Fig. 3D). These data indicate that GA transport by NPF2.12 and NPF2.13 is likely proton-coupled.

Proton-coupled transport is electroneutral if the proton-to-substrate stoichiometry is 1:1. To investigate whether the transport of GA by NPF2.12 and NPF2.13 is electrogenic, we used two-electrode voltage clamp electrophysiology to evaluate oocytes that express the transporters. Subtracting the currents elicited by oocytes at different membrane potentials in the absence of GAs from the currents elicited by oocytes in the presence of 500 µM GA₃ revealed that GA₃ transport by both NPF2.12 and NPF2.13 is associated with negative currents relative to control oocytes (Fig. 3E). The negative currents reflect a net positive influx of charges through both NPF2.12 and NPF2.13 upon GA₃ exposure. At low negative membrane potential (-20 mV), GA₃-induced currents in NPF2.12-expressing oocytes were of the same magnitude as those of control oocytes, whereas at high negative membrane potential (-120 mV), GA₃ induced-currents in NPF2.12-expressing oocytes were of the same magnitude as those of NPF2.13-expressing oocytes (Fig. 3E, F). This suggests that the apparent higher uptake of GA by NPF2.13 compared to NPF2.12 (Fig. 2B) likely reflects the membrane potential of resting Xenopus oocytes. In agreement with previous publications ^52,53, we detected nitrate import into oocytes that expressed NPF2.12 or NPF2.13, but this transport did not interfere with the GA transport capabilities as GA accumulation was not affected in GA/nitrate competition assays (Sup. Fig. 7B).

To elucidate whether these transporters are part of the GA transport mechanisms in the plant, we treated T-DNA knockout lines, which did not display any visible phenotypes (Sup. Fig. 8), with GA₄-Fl. npf2.12 and npf2.13 single mutant plants treated with GA₄-Fl displayed a significant reduction in accumulation in the endodermis compared to Col-0 plants (Fig. 3G), similar to reduced levels detected in npf3.1 mutants. To verify that the reduction in GA-Fl accumulation was not due to an off-target mutation, we repeated the test with an additional npf2.12 T-DNA insertion line (npf2.12-2) and obtained similar results (Sup. Fig. 9A). In addition, homozygous pNPF2.13:NPF2.13-YFP plants completely rescued the root GA₃-Fl phenotype (Sup. Fig. 9B).

We next generated plants ectopically expressing NPF2.12 and NPF2.13 fused to YFP (p35S:NPF2.12-YFP and p35S:NPF2.13-YFP, respectively) and treated them with GA₃-Fl. These lines showed a remarkably strong accumulation of GA₃-Fl in all root cells (Fig. 3H), supporting the hypothesis that NPF2.12 and NPF2.13 are GA transporters in planta. In order to test whether NPF2.12 and NPF2.13 have a dual specificity function and can also import ABA, we treated plants with fluorescently tagged ABA (ABA-Fl). ABA-Fl is non-bioactive but can be utilized to estimate ABA movement in the plant ³³. Overexpression of NPF2.12 or of NPF2.13 resulted in extreme ABA-Fl accumulation compared to Col-0 plants (Fig. 3H), implying that both transporters import GA and ABA in planta.

In order to address the subcellular localization of NPF2.12 and NPF2.13, we imaged the root epidermis cells of p35S:NPF2.12-YFP and p35S:NPF2.13-YFP lines. Both NPF2.12 and NPF2.13 are localized to the plasma membrane (Fig. 3I). Together, the in planta and oocyte results indicate that NPF2.12 and NPF.13 are dual-substrate, plasma membrane-localized GA and ABA importers.

NPF2.12 and NPF2.13 regulate root suberization

To further elucidate the biological function of NPF2.12 and NPF2.13, we generated NLS-YFP and GUS reporter lines to map NPF2.12 and NPF2.13 expression patterns. Confocal microscopy of plants expressing NLS-YFP driven by NPF2.12 and NPF2.13 native promoters revealed that in the root, NPF2.12 was expressed in the pericycle of the whole root and subsequently in the periderm of mature plants; NPF2.13, on the other hand, was expressed only in the shoot (Fig. 4A, Sup. Fig. 10). Analysis of pNPF2.12:GUS and pNPF2.13:GUS lines showed that the two transporters are expressed in the shoot vasculature (Fig. 4B). The fact that the NPF2.13 translational fusion construct (pNPF2.13:NPF2.13-Venus) introduced into the npf2.13 mutant background rescued the root GA₃-Fl accumulation phenotype indicated that the promoter region we cloned is sufficient and that NPF2.13 expression is restricted to the shoot (Sup. Fig. 9B).

Similar to npf2.14 mutants, npf2.12 and npf2.13 mutant plants displayed a reduction in suberization. Mutant roots stained with Nile red or Fluorol yellow showed a weaker fluorescence intensity than detected in Col-0 plants (Fig. 4C, Sup. Fig. 11). Similar results were obtained using additional mutant alleles for npf2.12 and npf2.13 (Sup. Fig. 12). The phenotype was enhanced in the npf2.12 npf2.13 double mutant line (Fig. 4C). Notably, GA₃ or ABA treatment completely rescued npf2.12 low-suberin phenotype. The phenotypes of npf2.13 and of the npf2.12 npf2.13 double mutant were completely rescued by ABA and largely rescued by GA₃(Fig. 4C). Quantification of suberin monomer content revealed that both npf2.12 and npf2.13 mutant roots accumulated ~40% less total suberin contents compared to the Col-0 attributed to lower levels of vanillic and ferulic acids, C22 fatty acid, C20 fatty alcohol, and C18:1(9), C22 and C26 ꞷ-hydroxyacids (Fig. 4D).

NPF2.12 and NPF2.13 regulate shoot-to-root GA translocation

Our previous work showed that GA₁₂, though not bioactive, is the primary GA form transported over long distances through the vasculature in Arabidopsis thaliana ¹². GA₁₂ can move through the xylem in a root-to-shoot manner and in the phloem in a shoot-to-root direction to regulate adaptive plant growth ^12,13. However, the mechanism regulating this process remains unknown ²¹. NPF2.13 was expressed strictly in the shoot (Fig. 4A, B), yet the knockout led to a phenotype in the root endodermis (Fig. 4C). This led us to hypothesize that the transporters are involved in the long-distance shoot-to-root translocation of GA. To test whether NPF2.12 and NPF2.13 facilitate shoot-born GA loading into the phloem, we quantified GAs content in phloem exudates collected from leaf petioles. The double npf2.12 npf2.13 loss-of-function mutant showed a striking reduction in GA₁₂ content in the collected phloem exudates (Fig. 5A). Other GA metabolites (GA₁₅, GA₂₄, GA₉, GA₄, GA₃₄, and GA₅₁) were not significantly reduced (Fig. 5A). ABA levels showed a mild decrease, significant when compared to Col-0 using two-tailed t-tests but not significant when using Dunnett’s multiple comparisons test (p ≤ 0.05). The results imply that NPF2.12 and NPF2.13 regulate GA, and possibly ABA, loading into the shoot phloem. In agreement, quantification of active GA₄content in the root_,which is downstream to GA₁₂ in the biosynthesis pathway, showed a significant reduction in the npf12 and npf13 single and double mutant lines (Fig. 5B). ABA content was also significantly lower in mutant roots compared to Col-0 (Fig. 5B).

To further test this hypothesis, we examined the expression pattern of the two transporters using reporter lines. In mature rosette, NPF2.12 and NPF2.13 were expressed in around the shoot apex and the main vascular vain (Fig. 5C). Cross-sections of pNPF2.12:GUS and pNPF2.13:GUS leaf petioles showed that both genes were expressed in the phloem companion cells (Fig. 5C). Next, we investigated whether these transporters are involved in long-distance GA movement from the shoot to the root. For this purpose, 16-day-old plants were grown on paclobutrazol, a GA biosynthesis inhibitor, for 4 days to deplete the plants of native GA, and GA₁₂was applied to a single leaf. We then quantified the DELLA protein REPRESSOR OF GA1-3 (RGA) abundance in the root. DELLA proteins act as repressors of the GA signal and are degraded in the presence of GA ⁵⁴. Time-course experiments in Col-0 plants showed a reduction in RGA accumulation in the root after GA₁₂ treatment, indicating GA₁₂ movement from the shoot to the root. On the other hand, in the npf2.12 npf2.13 double mutant there was no RGA degradation (Fig. 5D), signifying a reduced GA accumulation in the root.

To understand how the NPF2.12 and NPF2.13 mediated long-distance GA transport influences root growth, we performed a series of micrograftings between Col-0, npf2.12 npf2.13 and GA-deficient ga1-3 mutants and quantified root elongation rate. As expected, ga1-3 self-grafts had a significantly lower root elongation rate than the Col-0 self-grafts. npf2.12 npf2.13 self-grafts and Col-0 shoots grafted to ga1-3 roots were indistinguishable from the Col-0 self-grafts (Fig. 5E). The fact that a wild-type shoot could rescue the ga1-3 root elongation supports our hypothesis that GA can be transported from the shoot to the root to induce elongation. In comparison, npf2.12 npf2.13 shoots grafted to ga1-3 roots (ga1-3/npf2.12 npf2.13) showed a reduced root elongation compared to ga1-3/Col-0, indicating a partial requirement of NPF2.12 and NPF2.13 in long-distance GA transport (Fig. 5E). We hypothesize that once GA₁₂ is translocated to the roots, it can be converted to the bioactive GA₄ by the GA20ox and GA3ox enzymes, which are expressed in the root ⁵⁵. Profiling the expression pattern of the GA3ox promoters, catalyzing the last step in bioactive GA₄ hormone synthesis, showed that expression of these enzymes is restricted to the stele as previously reported ⁵⁵ (Fig. 5F). Together, these results imply that NPF2.12 and NPF2.13 function in GA₁₂ loading into the phloem for long-distance transport from the shoot to the root, with conversion to GA₄ taking place in the root stele.

Pericycle-specific vacuolar storage of GA and ABA at the phloem unloading zone facilitates endodermal suberin induction in the maturation zone.

To broaden our knowledge of GA and ABA distribution in the root and how it affects suberization, we created a mathematical model to simulate hormone distributions within the root cross-section, extending a modeling framework previously developed to study auxin transport ⁵⁶ to incorporate a vacuole compartment within each cell. We present the model results for GA; the transport and resulting distribution for ABA would follow a similar pattern. Using a multicellular template segmented from a root-cross-sectional image (Sup. Fig. 13), we incorporated into the model experimentally observed distributions of root GA transporters: NPF3.1 on endodermal cell membranes ^16,29, NPF2.12 on pericycle cell membranes (Fig. 3I, 4A), and NPF2.14 on the pericycle tonoplasts (Fig. 1K, 2A). The model simulated active GA transport via NPF3.1, NPF2.12, and NPF2.14, passive GA transport across both plasma membrane and tonoplast, GA synthesis and degradation, and GA diffusion within the apoplast with significantly reduced diffusion in the endodermal apoplast due to the presence of the Casparian strip.

To parameterize the model, permeabilities associated with each passive and active transport component were estimated using the oocyte data, and transport rates were then specified based on established pH and membrane potential values for plant cells ^56–58 (Sup. Table 4). An important factor is the source of the hormone. ABA2 and AAO3, which encode enzymes necessary for ABA biosynthesis, were previously shown to be expressed in the vasculature ⁵⁹. Bioactive GA₄ is also synthesized at high levels in the Arabidopsis stele ⁵⁵ (Fig. 5F). Considering these data, together with the phloem unloading zone ⁴¹, the docking belt for the long-distance shoot-to-root transported hormones, led us to specify the stele as the source of active GA and ABA in the model. With these assumptions and parameterization, the model predicted the wild-type distribution of both cytoplasmic and vacuolar hormone concentrations for the 5-day-old plants used in the experiments (Fig. 6A). These predictions revealed high levels of both cytoplasmic and vacuolar GA in the endodermis, where suberization later occurs. Mutations in npf3.1 and npf2.12 were predicted to lead to a reduction in endodermal GA cytoplasmic concentrations (Fig. 6A), in agreement with the loss of endodermal GA-Fl in these mutants (Fig. 3G) ¹⁶ and explaining their loss of suberization (Fig. 4C). Furthermore, while our experimental data suggested that NPF2.13 is not expressed in the root (Fig. 4A), the phloem sap and whole root GAs measurements suggest that NPF2.13 is an important contributor to the long-distance translocation of GA₁₂(Fig. 6A). The effects of the npf2.13 mutation can be simulated by reducing the stele-specific synthesis rate. When the synthesis rate was reduced to represent mutation of npf2.13, the model predicted a reduction in GA concentrations throughout the root cross-section, again providing an explanation of the reduction in observed suberization (Fig. 4C). In contrast, simulation of the npf2.14 mutation led to endodermal cytoplasmic GA concentrations that are higher than those in the wild-type, in agreement with the GA-Fl accumulation at the elongation zone (Fig. 1I). Why npf2.14 exhibits reduced suberization when endodermal accumulation is higher remained unclear. However, predicted vacuolar concentrations in the npf2.14 pericycle were much lower than in wild-type (Fig. 6A), leading us to hypothesize that NPF2.14 regulates GA levels both inside and outside of vacuoles in order to provide a GA source in the maturation zone, where GA is required for suberization. To test this hypothesis, we simulated the GA dynamics after reducing the synthesis rate to zero (mimicking cells leaving the phloem unloading zone) and found that the predicted endodermal cytoplasmic concentrations are maintained at higher levels with NPF2.14 present as cells mature (Fig. 6B). This suggests that the tonoplast pericycle-localized NPF2.14 ensures that GA concentrations are at levels necessary for mediation of suberization. Therefore, based on the model, we propose a pericycle-specific slow-release GA and ABA mechanism that explains how the two hormones are loaded into the pericycle vacuoles at the phloem unloading zone ⁴¹ and released from these vacuoles later on when the cells are mature (Fig. 6B). In this mechanism, GA and ABA unloaded from the phloem are transported into the pericycle and loaded into the vacuole to form a storage pool. When these cells reach the maturation zone, the GA and ABA that were stored in the pericycle vacuoles are transported by NPF3 into the endodermis to induce suberization.

In conclusion, the mathematical model confirmed that GA levels in the endodermis in wild-type roots were considerably higher than those in the npf3.1, npf2.12, and npf2.13 roots. Furthermore, the tonoplast-localized NPF2.14 ensures that there is a vacuolar source of GA that enables the cross-section to maintain high endodermal GA levels after cells leave the phloem unloading zone ⁴¹. The model predictions provide explanations for the suberization phenotypes observed in NPF mutants and emphasize that the pericycle is an NPF-mediated buffer-zone enabling cell-to-cell hormone movement from stele to endodermis.

In this work, we identified NPF2.14, a previously uncharacterized transporter, as a dual-specificity GA and ABA vacuolar transporter. To the best of our knowledge, NPF2.14 is the first known sub-cellular GA/ABA transporter. We showed that NPF2.14 is expressed in the pericycle to facilitate endodermal root suberization. Oocyte experiments showing that NPF2.14 exports GA from the cytosol, combined with NPF2.14’s localization to the tonoplast, indicates that NPF2.14 transports GA and ABA from the cytosol into the vacuole.

The results presented here suggest that the pericycle serves as a buffer zone, regulating the transitions of hormones from the stele to the endodermis (Fig. 6C). The stele acts as the source of bioactive GA and ABA in the root ^55,59. Both hormones have been shown to accumulate and affect their respective response specifically in the endodermis ^25,28,60,61. In addition, it appears that the GA and ABA do not simply flow through the pericycle but rather are loaded into the vacuole by NPF2.14 to form a reservoir for later developmental stages. We propose that the high levels of GA and ABA present in the phloem unloading zone are taken into the pericycle by NPF2.12. Once in the pericycle cells, NPF2.14 facilitates their import into the vacuole for storage. We speculate that a slow-release mechanism feeds the differentiating cells with GA and ABA, thus allowing suberin formation in the mature root (Fig. 6C). Vacuoles have been proposed to act as storage, modification, or degradation compartments for plant hormones ⁶². It is possible that tonoplast-localized NPF2.14 mediates the hormonal homeostasis balance that is needed for the proper execution of the developmental plan in the neighboring cell file. At this point, it is not clear if GA and ABA are stored at their bioactive form in the vacuole.

Confocal imaging of the NLS-YFP lines indicated that both NPF2.12 and NPF2.14 are expressed in the pericycle, mainly in the phloem poles (Fig. 2A, 4A). It is therefore possible that the hormone uptake is not carried uniformly throughout the pericycle ring, but rather amplified at the pericycle phloem poles cells. If so, do the two hormones retain polar distribution in the pericycle and in the subsequent endodermis layer? Or do the hormones have the ability to move within the two cell file rings? Future mathematical models and genetic work is required to address these questions.

The significantly reduced GA₁₂ content in npf2.12 npf2.13 double knockout phloem extracts implies that these transporters are required for GA₁₂ loading into the phloem and translocation of GA₁₂ from the shoot to the root. Thus, we hypothesize that NPF2.12 and NPF2.13, which are plasma membrane-localized importers expressed in the shoot phloem companion cells, are part of the long-sought mediators of long-distance GA shoot-to root translocation. Our results are in agreement with the previous finding that GA₁₂ is the main form of GA that is transported long distances through the plant ¹². In addition, we showed that GA3ox1 and GA3ox2, which catalyze the final step of active GA production, are expressed in the root stele ⁵⁵ (Fig. 5F). Thus, once GA₁₂ docks at the root stele, it is converted by GA3ox1 and GA3ox2 into the bioactive GA₄ form, which is then delivered from the stele to the endodermis by NPF2.12 and NPF3.1 (Fig. 6C).

It is intriguing that NPF2.12 and NPF2.13 act as GA₁₂ shoot-to-root transporters and also promote the delivery of the bioactive forms of GA and ABA to the endodermis. In both cases, a plasma membrane import activity is involved, but the substrate specificity differs. Since we showed that NPF2.12 and NPF2.13 are able to transport both intermediates of- and the bioactive forms of GA (Fig. 3B, Sup. Figure 7) and that GA₁₂ is present at high levels in the shoot phloem (Fig. 5A), we speculate that GA₁₂ is the primary substrate of the transporters. In the root stele, NPF2.12 recognizes bioactive ABA and GA₄, which are present in high concentrations due to being synthesized there ⁵⁹.

The presented work showed for the first time that GA promotes suberization; ABA is known to participate in this process ⁴⁹. This explains the physiological importance of GA accumulation in the endodermis. At this stage, it is not clear whether GA promotes endodermis suberization directly by, for example, direct binding and activation of the suberin biosynthesis factor by DELLA or DELLA co-partners, or whether suberization is a secondary effect of maturation or signaling through the ABA components. GA and ABA are antagonistic hormones ⁶³; however, our data revealed that the two hormones can act non-antagonistically to promote root suberization. The results are in agreement with the growth defects displayed by biosynthesis mutants of these hormones, which result in small dark green plants ^64–66. Thus, it could be that the two hormones act synergistically to promote growth and other processes under normal or stress conditions.

Plant material and growth conditions

All Arabidopsis thaliana lines are in a Col-0 background. T-DNA insertion lines were supplied by the Arabidopsis Biological Resource Center. PCR genotyping for homozygous lines was performed using the primers listed in Sup. Table 1. To generate npf2.12 npf2.13 double mutant, npf2.12 (SALK_138987) was crossed to npf2.13 (SALK_022429) to obtain an F1 population. F3 homozygous plant was selected by PCR genotyping.

Plants were sown on vertical plates containing 0.5 × Murashige-Skoog (MS) medium, 1% sucrose, and 0.8% agar, pH 5.7, stratified for 2 days at 4°C in the dark, then transferred to growth chambers (Percival CU41L5) at 21°C, 100 µE m^− 2 s^− 1 light intensity under long day light (16 h light/8 h dark). All plants in suberin quantification experiments were grown on 0.5 × MS medium, 1% sucrose, and 0.8% agar, pH 5.7 for 3 days and subsequently moved to 0.5 × MS medium without sucrose supplementation due to phenotype masking by the sucrose treatment. For low-nitrate experiments, plants were sown on MS with vitamins, nitrate free (Caisson labs MSP07-50LT), which was supplemented with 0.01 mM (Low nitrate) or 10 mM (High nitrate) KNO₃.

Hypocotyl cross sections

Sectioning and clearing were performed as describe in Ursache et al ⁴⁴. 3-Week-old hypocotyls were fixed in 4% PFA for an hour, rinsed twice in 1xPBS embedded in 5% agarose and sectioned to 150 µM slices using a Leica VT1000S vibratome. Slices were cleared using a ClearSee solution for 5 days. Following clearing sections were counterstained with 0.1% Calcofluor White in ClearSee solution for 30 min. Next, the seedlings were washed in ClearSee for 30 min with gentle shaking. For imaging, sections were mounted directly in ClearSee and imaged using a Zeiss LSM 780 inverted microscope.

Hormone application

Hormone was added to the agar medium at concentrations indicated in the figure legends. Seedlings were either germinated on media or moved after germination to treatment plates. GA-Fl (5 μM) was applied in liquid MS media for 16 h prior to imaging. For ga1 experiments, both Col-0 and ga1 seeds were imbibed in sterile water containing 5 μM GA₃ for 16 h to induce uniform germination. Following imbibition, seeds were washed three times in sterile water to wash away excess GA and were sown on MS plates.

Cloning of NPFs overexpression and reporter lines

NPF2.12 and NPF2.14 coding sequences were synthesized by Bio Basic Inc., cloned into pENTR/D-TOPO (Invitrogen K2400), and subsequently cloned into the pH7YWG2 destination vectors using the LR Gateway reaction (Invitrogen 11791). NPF2.12 and NPF2.14 promoters were amplified with the primers listed in Sup. Table 2 using a Phusion high-fidelity polymerase (New England Biolabs), cloned into pENTR/D-TOPO, and then cloned into pMDC7 vector for NLS-YFP reporters and pGWB3 vector for GUS reporters.

To generate pNPF2.13:GUS reporter, the promoter of NPF2.13 (1.7-kb fragment) was PCR amplified from Col-0 genomic DNA with appropriate primers listed in Sup. Table 2 and inserted into pDONR221 (Invitrogen) by Gateway cloning and recombined with pGWB633 ⁶⁷.

Imaging and analysis

Seedlings were stained in 10 mg/L^-1 propidium iodide (PI) for 5 min, rinsed, and mounted in water. Seedlings were imaged using a laser scanning confocal microscope (Zeiss LSM 780 inverted microscope), with argon laser set at 488 nm for fluorescein, 514 nm for YFP, and 561 nm for PI excitation. Emission filters used were 493-548 nm for fluorescein derivatives, 508-570 nm for YFP, and 583-718 nm for PI emission. Image analysis and signal quantification were done with the measurement function of ZEN lite software. The number of quantified biological repeats and sampling points is indicated for each graph in figure legends.

Root length characterization

For root length measurements, seedlings were imaged using Zeiss Stemi 2000-C stereo microscope and measured using ImageJ software (http://rsbweb.nih.gov/ij/index.html).

Histochemical GUS staining

Plants were immersed in 100 mM sodium phosphate buffer (pH 7.0) containing 0.1% Triton X-100, 1 mM 5-bromo-4-chloro-3-indolyl-β-D-glucuronic acid cyclohexylammonium salt (Sigma-Aldrich), 2 mM potassium ferricyanide, and 2 mM potassium ferrocyanide. Plants subject to vacuum treatment for 10 min and then incubated at 37 °C for 16 h. Tissues were cleared with 30%, 50%, and 70% ethanol for 30 min in each concentration and imaged using an AxioZoom 16, Zeiss binocular microscope.

For cross-sectioning of GUS-stained leaf petioles, after clearing in 70% ethanol, the samples were fixed in FAA solution (3.2% formaldehyde, 5% acetic acid, 50% ethanol) for 30 min and kept overnight at 4°C. The samples were then dehydrated in an ethanol gradient ranging from 50% to 96%, and incubated in 2% eosin overnight at 4° C. After several washes in 96% ethanol, the samples were progressively rehydrated in ethanol/HISTO-CLEAR II (Electron Microscopy Sciences) solution, incubated in 50% HISTO-CLEAR II 50% PARAPLAST PLUS (McCormick Scientific) at 60°C for 2 h, and embedded in 100% PARAPLAST PLUS. Paraffin-embedded samples were cross-sectioned with LEICA RM2155 microtome and imaged using a Leica Leitz Dmrb microscope.

Nile red suberin staining, imaging and quantification.

Nile red suberin staining was performed as described by Ursache et al. ⁴⁴. In short, 5-day-old seedlings were fixed in paraformaldehyde for 1 h under gentle agitation and washed twice in phosphate-buffered saline, pH 7.4. Plants were covered in filtered 0.05% Nile red (Acros Organics, 7385-67-3) solution dissolved in ClearSee for 16 h. Following staining, plants were washed three times in ClearSee for 30 min each wash. Next, plants were counterstained with 0.1% calcofluor white (Glentham Life Sciences, 4404-43-7) dissolved in ClearSee for cell wall imaging. After 30 min, plants were washed in ClearSee for 30 min. Plants were mounted directly in ClearSee on slides and imaged with a Zeiss LSM780 confocal microscope. Images were taken from the upper part of the root and under the root-hypocotyl junction with an argon laser set at 514 nm for Nile red excitation and 405 nm for calcofluor excitation. Emission filters used were 561-753 nm filter for Nile red and 410-511 nm filter for calcofluor emission. Fluorescence intensity was assessed using the Zen software from 5 endodermal cells per root.

Phylogenetic tree

Protein sequences for Arabidopsis thaliana NPF family members were retrieved from TAIR (https://www.arabidopsis.org). Phylogenetic relationships were defined using Phylogeny.fr (http://www.phylogeny.fr/) and visualized with FigTree software (http://tree.bio.ed.ac.uk/software/figtree/).

Transport assays in Xenopus oocytes

Coding sequences were cloned into the pNB1u vector, and complementary RNA (cRNA) was produced as described in Wulff et al. ³⁴. Xenopus oocyte assays were performed as described previously ³⁴. Defolliculated Xenopus laevis oocytes (stage V-VI) were purchased from Ecocyte Biosciences and were injected with 25 ng cRNA in 50.6 nl using a Drummond Nanoject II and incubated for 2-4 days at 16 °C in HEPES-based kulori (90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM HEPES, pH 7.4) before use. Oocytes were pre-incubated in MES-based kulori (90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM MES, pH 5) for 4 min and were then transferred to phytohormone-containing MES-based kulori for 60 min. After washing three times in 25 ml HEPES-based kulori followed by one wash in 25 ml deionized water, oocytes were homogenized in 50% methanol and stored for >30 min at -20 °C. Following centrifugation (25000 g for 10 min 4 °C), the supernatant was mixed with deionized water to a final methanol concentration of 20% and filtered through a 0.22-µm filter (MSGVN2250, Merck Millipore) before analytical LC-MS/MS as described below. For nitrate assays, sodium chloride in kulori was substituted for equimolar sodium nitrate in order not to affect the membrane potential.

Quantification of phytohormone content by LC-MS/MS

Compounds in the diluted oocyte extracts were directly analyzed by LC-MS/MS. The analysis was performed with modifications from the method described in Tal et al. ¹⁶. In brief, chromatography was performed on an Advance UHPLC system (Bruker). Separation was achieved on a Phenomenex Kinetex 1.7u XB-C18 column (100 x 2.1 mm, 1.7 µm, 100 Å) with 0.05% v/v formic acid in water as mobile phase A and acetonitrile with 0.05% formic acid (v/v) as mobile phase B. The gradients used for elution of GAs were 0-0.5 min, 2% B; 0.5-1.3 min, 2-30% B; 1.3-2.2 min 30-100% B, 2.2-2.8 min 100% B; 2.8-2.9 min 100-2% B; and 2.9-4.0 min 2% B. The gradients used for elution of ABA were 0-0.5 min, 2% B; 0.5-1.2 min, 2-30% B; 1.2-2.0 min, 30-100% B; 2.0-2.5 min, 100%; 2.5-2.6 min, 100-2% B; and 2.6-4.0 min, 2% B. The mobile phase flow rate was 400 µl min^-1, and column temperature was maintained at 40 °C. The liquid chromatography was coupled to an EVOQ Elite triple quadrupole mass spectrometer (Bruker) equipped with an electrospray ion source operated in positive and negative ionization mode. Instrument parameters were optimized by infusion experiments with pure standards. For analysis of GAs, the ion spray voltage was maintained at +4000 V and -4000 V in positive and negative ionization mode, respectively, and the heated probe temperature was set to 200 °C with probe gas flow at 50 psi. For ABA, the ion spray voltage was maintained at -3300 V in negative ionization mode, and heated probe temperature was set to 120 °C with probe gas flow at 40 psi. Remaining settings were identical for all analytical methods with cone temperature set to 350 °C and cone gas to 20 psi. Nebulizing gas was set to 60 psi and collision gas to 1.6 mTorr. Nitrogen was used as probe and nebulizing gas, and argon as collision gas. Active exhaust was constantly on. Multiple reaction monitoring was used to monitor analyte parent ion to product ion transitions for all analytes. Multiple reaction monitoring transitions and collision energies were optimized by direct infusion experiments. Detailed values for mass transitions can be found in Supplemental Table 3. Both Q1 and Q3 quadrupoles were maintained at unit resolution. Bruker MS Workstation software (Version 8.2.1) was used for data acquisition and processing. Linearity in ionization efficiencies were verified by analyzing dilution series of standard mixtures. Sinigrin glucosinolate was used as internal standard for normalization but not for quantification. Quantification of all compounds was achieved by external standard curves diluted with the same matrix as the actual samples. All GAs were analyzed together in a single method. GA₁₂ suffered from severe ion suppression when combined with the other GAs in the standard curve, thus quantification was not achieved for GA₁₂.

Root suberin monomer profiling by GC-MS

Suberin monomers were extracted from Col-0 and mutant roots according to the protocols previously described by ^68,69. A sample volume of 1 µL was injected in splitless mode on a GC-MS system (Agilent 7693A Liquid Auto injector, 8860 gas chromatograph, and 5977B mass spectrometer). GC was performed (HP-5MS UI column; 30 m length, 0.250 mm diameter, and 0.25 µm film thickness; Agilent J&W GC Columns) with injection temperature of 270°C, interface set to 250°C, and the ion source to 200°C. Helium was used as the carrier gas at a constant flow rate of 1.2 mL min^-1. The temperature program was 0.5 min isothermal at 70°C, followed by a 30°C min^-1 oven temperature ramp to 210°C and a 5°C min^-1 ramp to 330°C, then kept constant during 21 min. Mass spectra were recorded with an m/z 40 to 850 scanning range. Chromatograms and mass spectra were evaluated using the MSD ChemStation software (Agilent). Integrated peaks of mass fragments were normalized for sample dry weight and the respective C32 alkane internal standard signal. For identification, the corresponding mass spectra and retention time indices were compared with the NIST20 library as well as in-house spectral libraries.

Xenopus oocyte injection based efflux transport assays

For injection-based export assays, on the second day of gene expression, oocytes were injected with 23 nl 8.2 mM in 98 mM KCl, 1 mM CaCl₂, 10 mM HEPES, pH 7.4. T1 oocytes were left 10 min to heal and were then transport was evaluated as described above. T2 oocytes were left for approximately 20 h in HEPES-based kulori at 16° C, followed by transport analysis.

Quantification of nitrate from oocytes by HPLC

Nitrate concentration in the oocyte extracts was quantified using a Dionex ICS-2100 anion exchange chromatography system (Thermo Scientific). The separation was done on a Dionex IonPac AG11-HC analytical column coupled to the AS11-HC guard column (Thermo Scientific). The columns were connected to a Dionex AERS 500 anion suppressor (Thermo Scientific). The analyses were performed under the following conditions: sample injection volume 4.8 µl, column temperature 30 °C, flow rate of 0.38 ml/min, isocratic eluent gradient using 30 mM KOH solution in QH₂O, suppressor current of 29 mA, and runtime of 15 min. The nitrate detection was done at 220 nm using a Dionex UltiMate 3000 (Thermo Scientific). QH₂O water dilutions of Dionex Combined Seven Anion Standard (Thermo Scientific) were used to create a standard calibration curve. Accuracy and precision of the quantification was checked by including samples of potassium nitrate throughout the sequence.

pH measurements of oocyte lumen

The pH stabilization was performed as described previously ³⁴. pH-electrodes were pulled from borosilicate glass capillaries (KWIK-FIL TW F120-3 with filament) on a vertical puller (Narishige Scientific Instrument Lab), baked for 120 min at 220 °C and silanized for 60 min with dimethyldichlorosilane (Silanization Solution I, Sigma Aldrich). Electrodes were backfilled with a buffer containing 40 mM KH₂PO₄, 23 mM NaOH and 150 mM NaCl (pH 7.5). The electrode tip was filled with a proton-selective ionophore cocktail (hydrogen ionophore I cocktail A, Sigma-Aldrich) by dipping the tip into the cocktail. Oocytes, as described above, were placed in freshly made HEPES-based ekulori (2 mM LaCl₃, 90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM HEPES pH 7.4) for at least 30 min prior to three-electrode voltage clamp experiments. Before each oocyte a pH calibration curve was made for each oocyte using 100 mM KCl pH 5.5, 100 mM KCl pH 6.5 and 100 mM KCl pH 7.5. Oocytes were clamped at 0 mV and perfused with HEPES-based ekulori pH 7.4, followed by MES-based ekulori (2 mM LaCl₃, 90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM MES pH 5) and internal pH response was measured continuously as a function of external pH change.

Membrane potential measurements

Membrane potentials of oocytes were measured using the automated two-electrode voltage clamp system, Roboocyte2 (Multi channel systems), in ekulori (90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM MES, 2 mM LaCl₃, pH 5) with electrodes backfilled with 1 M KCl and 1.5 M potassium acetate. All oocytes were measured using the same electrodes with a resistance of 280-350 kΩ. The experiment was terminated when the resistance of one of the electrodes shifted to approximately 600 kΩ.

Two-electrode voltage clamp electrophysiology

The electric signal elicited by GA treatment of oocytes was measured using the automated two-electrode voltage clamp system Roboocyte2 (Multi channel systems), in ekulori (90 mM NaCl, 1 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, 5 mM MES, 2 mM LaCl3 pH 4.5 or pH 5) with electrodes (resistance 280-1000 kΩ) backfilled with 1 M KCl and 1.5 M potassium acetate. Oocytes were clamped at – 60 mV, and IV curves were obtained before and after substrate addition. Substrate dependent currents were calculated by subtracting currents before addition of substrate from currents after addition of substrate.

Root hormone quantification

Hormone extraction and analysis was performed as described in Zhang et al., 2021. Standards (both labelled and non-labelled) were obtained from Olchemim Ltd. (Olomouc, Czech Republic) and National Research Council (NRC-CNRC, Canada). Standard grade solvents were used for sample preparation, Methanol, Acetic acid (LiChrosolv, Sigma-Aldrich, USA), Acetonitrile (J.T.Baker, Avantor, PA, USA), Formic acid (Honeywell Fluka, Thermo Fisher Scientific, MA, USA) and de-ionized water (Milli-Q, Synergy-UV millipore system, USA). Briefly, root tissue frozen in liquid nitrogen was grounded using motor and pestle. Around 200 mg of root sample was measured from ground powder and extracted with ice cold methanol/water/formic acid (15/4/1 v/v/v) added with deuterium labelled internal standards (IS). Similar concentrations of IS of abscisic acid and gibberellin (GA₄) were added into samples and calibration standards. The samples were purified using Oasis MCX SPE cartridges (Waters, USA) according to manufacturer’s protocol. The samples were injected on Acquity UPLC BEH C18 column (1.7 µm, 2.1x100 mm, Waters; with gradients of 0.1% acetic acid in water or acetonitrile), connected to Acquity UPLC H class system (with Waters Acquity QSM, FNR sample manager and PDA) coupled with UPLC-ESI-MS/MS triple quadrupole mass spectrometer (Xevo TQ-S, Waters, equipped with ESI probe) for identification and quantification of hormones. The hormones were measured using MS detector, both in positive and negative mode, with two MRM transitions for each compound. External calibration curves were constructed with hormone standards added with IS, used for quantification, and calculated through Target Lynx (v4.1; Waters) software by comparing the ratios of MRM peak areas of analyte to that of internal standard.

Phloem extract and hormone quantification

Rosette leaves of 5-week-old Col-0 and npf2-12 npf2-13 mutant plants (before bolting) were cut with a razor blade at the base of the petiole, and each leaf was dipped in a tube containing 80 µL of exudation buffer (50 mM potassium phosphate buffer, pH7.6, 10 mM EDTA). Exudation was carried out for 3h in dark in high humidity to limit transpiration. Exudation of 75 leaves was regrouped and concentrated under vacuum centrifugation. Hormone contents in phloem exudates were determined by UPLC system-MS/MS (Waters Quattro Premier XE). Concentrated residue of phloem sap was resuspended with 80% methanol-1% acetic acid including 17-²H₂-labeled GA internal standards (Olchemim), mixed and passed through an Oasis HLB column_. The dried eluate was dissolved in 5% acetonitrile-1% acetic acid, and the GAs were separated by UPHL chromatography (Accucore RP-MS column 2.6 µm, 100 x 2.1 mm; ThermoFisher Scientific) with a 5 to 50% acetonitrile gradient containing 0.05% acetic acid, at 400 µL/min over 22 min. The concentrations of GAs in the extracts were analyzed with a Q-Exactive mass spectrometer (Orbitrap detector; ThermoFisher Scientific) by targeted SIM using embedded calibration curves and the Xcalibur 2.2 SP1 build 48 and TraceFinder programs.

Grafting assays.

Grafting was performed without collars on water imbibed 0.45 µM MCE membrane (Millipore) between hypocotyls of rootstocks and scions of 6-day-old seedlings grown on 1x MS agar plate. Grafted seedlings were then kept vertically to recover, for 5 days under constant humidity. Successful grafts were transferred onto ½x MS agar plates and grown under a 16h photoperiod at 22°C. Root growth was measured every day for 4 days.

DELLA degradation assays.

12-day-old seedlings were transferred to 1x MS agar modified medium without nitrogen (bioWORLD plant media) supplemented with 0.5 mM KNO₃ and 1 µM paclobutrazol (Sigma). 4 days after transfer, a drop of GA₁₂ (5 µl at 1 µM) was placed on one of the first two leaved formed. Roots were collected 6, 12 and 24h after adding GA₁₂. Total proteins were extracted in 2x SDS-PAGE sample buffer and separated on 10% SDS-PAGE gel. After transfer onto membrane, immunoblots were performed using a 2000-fold dilution anti-RGA (Agrisera) and a 10000-fold dilution of peroxidase-conjugated goat anti-rabbit (Thermo Fisher Scientific). Signals were detected with Fusion FX (Vilber) using Immobilon Forte Western HRP Substrate (Millipore). The blot was subsequently stained with Coomassie blue. Quantification of the signals was determined using ImageJ package.

Mathematical model.

Root templates were segmented from an experimental image using the CellSeT image analysis tool ⁷⁰ (Sup. Fig. 13). We used CellSeT to manually assign a cell type to each cell and then read the geometrical and cell-type data into a tissue database (based on the OpenAlea tissue structure ⁷¹), extending the data structure to incorporate vacuolar compartments within each cell. The geometrical, topological and transporter-distribution data were used to form a system of ordinary differential equations (ODEs) to describe the GA transport, synthesis and degradation within the multicellular root cross-section. Parameters associated with the passive and transporter-mediated transport components were estimated using the oocyte data (Fig. 1A, Fig. 3B) and the remaining parameter values were obtained from the literature (Sup. Table 4). These ODEs were simulated using the solve_ivp package in python 3.6.5. Full details of the model equations and assumptions are provided as Supplementary text.

Acknowledgments

We thank Daria Binenbaum for the illustrations and Peter Hedden (Rothamsted Research) for sharing pGA3ox1-4:GUS seeds. Funding: This work was supported by grants from the Israel Science Foundation (2378/19 and 3419/20 to E.S.), the Human Frontier Science Program (HFSP—RGY0075/2015 and HFSP—LIY000540/2020 to E.S., H.H.N.-E. and L.R.B.), Danmarks Grundforskningsfond (DNRF99 to H.H.N.-E.), the European Research Council (757683-RobustHormoneTrans to E.S.), the Constantiner Travel Fellowship (to J.B.), the Centre National de la Recherche Scientifique (to L.S-A. J-M.D. and P.A.), the French Ministry of Research and Higher Education studentship (to L.C.).

Author contributions

J.B. performed the research and wrote the manuscript. N.W. performed the oocyte transporter assays. L.C., L.S-A., J-M.D. and P.A carried out long-distance transport assays. K.K. performed the mathematical modelling. I.T. assisted in cloning overexpression and reporter lines. H.V. and A.A. quantified root GA and ABA content. M.A. helped with genotyping T-DNA mutant lines. Y.Z. helped with npf mutant identification. D.R. and L.R. assisted in cross sectioning and staining. E.C. quantified hormone content in the phloem sap. E.M. and H.C. performed suberin monomer quantifications. S.L. and R.W. synthesized fluorescently tagged hormones. V.N. and C.C. helped with nitrate and hormones quantification in oocyte assays, respectively. L.B., P.A., H.H.N.-E. and ES designed and supervised the work and edited the manuscript. All authors discussed the results and commented on the manuscript.

Competing interests: The authors declare that they have no competing interests.

Data and materials availability: All the data supporting the findings of this study are available within the article and the Supplementary Materials.

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Gibberellin and abscisic acid transporters facilitate endodermal suberin formation in Arabidopsis

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NPF2.14 is a tonoplast-localized GA and ABA transporter

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