PIN2-mediated self-organizing transient auxin flow contributes to auxin maxima at the tip of Arabidopsis cotyledons

Pérez-Henríquez, Patricio; Nagawa, Shingo; Liu, Zhongchi; Pan, Xue; Michniewicz, Marta; Tang, Wenxin; Rasmussen, Carolyn; Cui, Xinping; Van Norman, Jaimie; Strader, Lucia; Yang, Zhenbiao

doi:10.1038/s41467-024-55480-8

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Published: 05 February 2025

PIN2-mediated self-organizing transient auxin flow contributes to auxin maxima at the tip of Arabidopsis cotyledons

Nature Communications volume 16, Article number: 1380 (2025) Cite this article

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Abstract

Directional auxin transport and formation of auxin maxima are critical for embryogenesis, organogenesis, pattern formation, and growth coordination in plants, but the mechanisms underpinning the initiation and establishment of these auxin dynamics are not fully understood. Here we show that a self-initiating and -terminating transient auxin flow along the marginal cells (MCs) contributes to the formation of an auxin maximum at the tip of Arabidopsis cotyledon that globally coordinates the interdigitation of puzzle-shaped pavement cells in the cotyledon epidermis. Prior to the interdigitation, indole butyric acid (IBA) is converted to indole acetic acid (IAA) to induce PIN2 accumulation and polarization in the marginal cells, leading to auxin flow toward and accumulation at the cotyledon tip. Once IAA levels at the cotyledon tip reaches a maximum, it activates pavement cell interdigitation as well as the accumulation of the IBA transporter TOB1 in MCs, which sequesters IBA to the vacuole and reduces IBA availability and IAA levels. The reduction of IAA levels results in PIN2 down-regulation and cessation of the auxin flow. Hence, our results elucidate a self-activating and self-terminating transient polar auxin transport system in cotyledons, contributing to the formation of localized auxin maxima that spatiotemporally coordinate pavement cell interdigitation.

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Introduction

Proper formation and morphogenesis of an organ requires not only the differentiation of cells and tissues, but also their coordinated growth in time and space, but the mechanisms for this coordination are mostly unexplored in multicellular organisms. The Arabidopsis cotyledon is an outstanding and trackable example of growth coordination and morphogenetic pattern formation towards the building of an efficient light-capturing flat structure. The expanding epidermal layers of pavement cells (PCs) on both sides of the flat structure accommodate the expanding mesophyll cells. On the edges, a file of elongated cells called the marginal cells (MCs) surrounds the cotyledon. During cotyledon growth, PCs undergo a unique multi-polar expansion going from near cuboidal cells towards interdigitated puzzle-shape cells¹. The PC interdigitation event occurs with specific auxin signaling coordination throughout the cotyledon². The small molecule phytohormone auxin acts as both a local and global signal to coordinate the PC interdigitation^3,4,5,6. Local auxin activates cell surface signaling cascades that are independent of auxin-induced gene transcription and directly leads to PC interdigitation by regulating ROP GTPase-dependent cytoskeletal reorganization^1,3,4,6,7. Auxin maxima and gradients peaking in a specific region of developing organs are a common feature in plants^8,9. It serves as guidance for proper patterning, growth regulation, cell type differentiation and organogenesis¹⁰. Auxin accumulation at the cotyledon’s tip is a widely reported observation¹¹, however, the mechanism for the generation of this tip-high auxin remains unknown.

At the heart of auxin maximum/gradient formation is the integrated regulation of various mechanisms impacting cellular auxin levels^12,13, including PIN-mediated polar auxin transport ^14,15,16, AUX1-mediated auxin influx¹⁷, ABC transporter mediated auxin efflux^18,19, auxin biosynthesis and metabolism^20,21,22 and local subcellular sequestration of auxin or its precursors^23,24,25. The absence of auxin transport and/or auxin biosynthesis can partially or completely disrupt the auxin gradients or maxima, proper patterning, and development. Thus, important developmental information is engraved in the spatial distribution of auxin^8,26,27,28, but the nature of this information and the mechanism by which it controls the formation of specific auxin gradients/maxima are still enigmatic.

Auxin is synthesized predominantly from tryptophan through the intermediate INDOLE 3-PYRUVIC ACID (IPyA) in the TAA/YUCCA-regulated IPyA pathway²⁹. Additionally, the INDOLE BUTYRIC ACID (IBA) pathway can locally boost auxin levels through the metabolic conversion of this auxin precursor into active auxin INDOLE ACETIC ACID (IAA). The IBA-to-IAA conversion pathway has been implicated in several developmental processes such as cotyledon expansion, organ size determination and lateral root formation²⁵. It occurs through several enzymatic reactions in the peroxisome where the function of enoyl CoA-hydratase 2, ECH2, is a limiting step²¹. Cotyledons of the ech2ibr1,3,10 quadruple mutant with impeded IBA-derived auxin exhibit severely impaired PC morphogenesis^2,30. Cytoplasmic IBA levels can also be locally reduced by TOB1 (TRANSPORT OF IBA 1)-mediated vacuole transport²⁴. Subcellular compartmentalization of IBA has been shown to regulate morphogenetic processes such as lateral root formation²⁴.

Despite our extensive knowledge on the critical contribution of both auxin biosynthesis and polar transport to the formation of auxin maxima, it is unclear how these processes are regulated and coordinated to generate and maintain local accumulation of auxin. Here we address this question by using the unique cotyledon system, in which the auxin maximum at the cotyledon’s tip is thought to guide its expansion. We showed that at the cotyledon edge, a file of elongated marginal cells accumulates polarly localized PIN2 forming a polar auxin transport system towards the tip. This auxin transport system is only transiently established via a self-activating and self-terminating mechanism to robustly coordinate PC interdigitation.

Results

PIN2-expressing margin cells coordinate pavement cell interdigitation

In developing organs, auxin maxima are formed via the polarized action of the plasma membrane (PM)-localized PIN-family auxin transporters^8,15. Published transcriptional data reveals that PINs are induced during germination. Among all PINs, PIN2 shows the strongest expression in early imbibed seeds. (Supplementary Fig. 1a). Interestingly, we found PIN2-GFP³¹ protein accumulated specifically in the lateral borders of early expanding cotyledons, forming a file of contiguous marginal cells (MCs) harboring PIN2 at their apical ends, resembling an “auxin transport pipeline” discharging at the cotyledon tip (Fig. 1a and Supplementary Fig. 2b). The strong signal at the cotyledon tip does not reflect true PIN2-GFP but autofluorescence, as confirmed by spectral scanning microscopy (Supplementary Fig. 2a–c). Although we cannot exclude the role of other PINs, the apical localization of PIN2-GFP hints a role for PIN2 transporting auxin toward the tip of cotyledons along their marginal cells.

**Fig. 1: PIN2 accumulation in MCs precedes and plays a role in coordinating PC morphogenesis.**

To further understand the correlation between PC interdigitation and PIN2-driven auxin transport, we analyzed the dynamics of PIN2-GFP accumulation in expanding cotyledons, from 12 to 96 Hours After Plating (HAP) (Fig. 1b). Strikingly, PIN2-GFP accumulation pattern in cotyledons was transient (Fig. 1c). Quantification revealed that PIN2 accumulation starts and consolidates before PC interdigitation is initiated (24-36 HAP) and is depleted once the interdigitation process is completed (72 HAP) (Fig. 1d). This suggests that PIN2-mediated auxin transport along MCs is involved in PC interdigitation presumably through contributing to the cotyledon’s tip auxin maxima.

Importantly, the auxin transcriptional response marker DR5::GUS exhibited greatly reduced GUS signal at the cotyledon tip in the PIN2 loss of function mutant eir1-1/pin2³², compared to wild type (Fig. 1e). This reduced GUS signal was corroborated by the quantification of GUS activity in a fluorometric assay (Fig. 1f). Additionally, pin2 showed less interdigitated and smaller PCs at the cotyledon tip (Fig. 1g, h and Supplementary Fig. 1c). Similar phenotypes were also observed in the insertional mutant allele pin2-T³³ (Supplementary Fig. 1d, e). These results imply that PIN2, which is specifically expressed in MCs, has a cell non-autonomous function in the promotion of PC interdigitation in cotyledons. To decipher the functional relevance of MCs in the regulation of PC interdigitation, we laser-ablated all MCs in a cotyledon prior to the initiation of PC interdigitation. At 24 h after ablation, the cotyledon displayed a strong inhibition of PC interdigitation, although its physiological status remained normal (Supplementary Fig. 2d–h). The laser ablation experiments hinted a role for PIN2-expressing MCs in the regulation of PC interdigitation. Altogether, low interdigitation in pin2 cotyledon’s tip, low auxin response in pin2 cotyledon tips, consistent apical (towards tip) localization of PIN2 in consecutive MCs, and the importance of PIN2-expressing MCs revealed by laser ablation, suggest the importance of PIN2-mediated auxin transport along the cotyledon margins towards the tip for the formation of auxin maxima to coordinate PC interdigitation.

If MCs are indeed transporting auxin, we should be able to detect increased auxin levels in these cells after PIN2 is accumulated. We tested this by comparing nuclear auxin marker before and after PIN2-GFP accumulation (Fig. 2a, 24 vs 36 HAP) in stable non-segregant lines containing both PIN2-GFP and the ratiometric auxin reporter R2D2³⁴. R2D2 reports auxin-induced degradation of DII-Venus normalized by an auxin non-responsive version mDII protein (Fig. 2b). The nuclear quantification of R2D2 ratiometric index along MCs showed a ~ 50% increment after PIN2-GFP accumulation started (Fig. 2c), indicative of increased auxin levels along MCs. Taken together we conclude that a transient PIN2-based auxin flow through the cotyledons margin contributes to the tip auxin maximum and plays a role in coordinating the PC interdigitation process.

**Fig. 2: Auxin promotes its own PIN2-mediated efflux along MCs.**

Auxin positively feedback regulates polar auxin transport along the MCs

The transient nature of PIN2 accumulation presented an intriguing question: What are the underlying mechanisms behind initiation and termination of the PIN2 accumulation along MCs? The increased auxin levels together with PIN2-GFP polarization is consistent with the well-known effect of auxin regulating PIN expression³⁵. Therefore, we supposed that auxin might positively regulate PIN2-mediated auxin transport along MCs. Indeed, we detected a dose-dependent rapid increase in the level of PIN2 localized to the apical plasma membrane after auxin treatments for 0.5 h, with a significant increase beginning at 100 nM IAA (Fig. 2d–f). PIN2 plasma membrane localization was also more polarized after auxin treatments (Supplementary Fig. 3d). Conversely, interruption of auxin transport by NPA treatment disrupted PIN2 polar localization (Supplementary Fig. 3a–c) and PC interdigitation (Supplementary Fig. 3e, f). Together these data suggest that an active “tip-wards” auxin transport along the MCs is initially organized by auxin itself.

IBA-derived auxin regulates PIN2 accumulation along MCs

To understand the biosynthetic origin of the auxin that flows along MCs, we explored the INDOLE 3-PYRUVIC ACID (IPyA) and INDOLE BUTYRIC ACID (IBA) pathways as possible sources of auxin. To this end, we used the presence or absence of key enzymes on each pathway as markers of their respective pathway activity. ECH2p::YFP-ECH2³⁰ and cytoplasmic TAAp::GFP-TAA1^11,36 were used to mark IBA and IPyA pathways, respectively. GFP-TAA1 does not accumulate in MCs prior to PIN2-GFP accumulation (Supplementary Fig. 4a), suggesting the IPyA pathway does not initially trigger PIN2 accumulation. Furthermore, transcripts coding for enzymes working in the IPyA pathway (TAA1, TAR2 and YUC1-11) are weakly or not expressed, while those for the IBA pathway (IBR1, IBR3, IBR10, ECH2) are highly expressed at this developmental stage (Supplementary Fig. 4c, d). However, GFP-TAA1 did accumulate in MCs and epidermal cells after PIN2-GFP was already accumulated (48 HAP), suggesting that the IPyA pathway may work in reinforcing auxin levels (Supplementary Fig. 4a, white arrows). In fact, chemical inhibition of the IPyA pathway reduced the accumulated PIN2 levels by ~25% but did not inhibit the formation of the PIN2-based ‘pipeline’ along MCs (Supplementary Fig. 4e, f).

In contrast, YFP-ECH2 showed clear accumulation in MCs prior to the PIN2 establishment (Fig. 3a) and a clear additional accumulation at the cotyledon tip after PIN2 establishment (Supplementary Fig. 4b). ECH2-labeled punctate structures could indicate peroxisomal localization (Fig. 3a, white arrows and intensity profiles), as shown previously in roots³⁰. In peroxisomes, IBA is converted to IAA³⁰. This punctate localization of ECH2 in MCs suggests the biosynthesis of IBA-derived IAA in those cells. Thus, we anticipated that PIN2 establishment along MCs would be compromised in IBA-to-IAA conversion mutants. Indeed, ech2ibr10 showed decreased levels of apical PIN2-GFP in cotyledon MCs at 48 HAP (Fig. 3b, c). Consistent with reduced auxin maxima, ech2ibr10 cotyledons were smaller and hyponastic (Supplementary Fig. 4g) and exhibited defects in PC interdigitation, as reported previously^2,30 (Fig. 3 d, e). Disrupted auxin transport through the cotyledon margins would explain the reduced DR5::GUS expression previously observed in the tip of ech2ibr10 cotyledons, whose severe PC interdigitation defects were rescued by exogenous auxin². Altogether, these results support the hypothesis that predominantly IBA-to-IAA conversion drives the onset of PIN2 accumulation along MCs, resulting in the formation of the tip auxin maximum that spatiotemporally coordinates PC interdigitation².

**Fig. 3: IBA-derived auxin regulates PIN2 accumulation along cotyledons margin cells.**

The distribution of TOB1 to the tonoplast is associated with PIN2 downregulation

The fact that the apical PIN2 levels were greatly reduced in the ech2ibr10 mutant prompted us to hypothesize that the PIN2 down-regulation observed around 60-72 HAP in wild type cotyledon MCs (Fig. 1d) may result from a reduction in the IBA-to-IAA conversion in those cells. One possible mechanism for this reduction would be through down-regulation of cytoplasmic IBA availability. Recently, TOB1 was described to reduce the IBA pool by importing IBA into vacuoles, consequently altering lateral root formation²⁴. Thus, TOB1 might also regulate IBA availability in cotyledon’s MCs, considering that TOB1 and PIN2 expression patterns are very similar in shoots, both being expressed in epidermal cells in a highly specific fashion (Supplementary Fig. 5a). In fact, the translational reporter TOB1p::YFP-TOB1 line (hereafter YFP-TOB1) showed highly specific TOB1 protein accumulation at the cotyledon’s MCs during the first 4 days of seedling growth, with some additional expression in neighboring cells (Supplementary Fig. 5c). Notably, the pattern of YFP-TOB1 subcellular distribution changes during cotyledon growth probably reflecting a change in the TOB1 function. YFP-TOB1 signal became reticular when PIN2 accumulation peaked (Fig. 4a, 48 HAP), whereas it was mainly vacuolar/prevacuolar (ring-like) when PIN2 disappeared (Fig. 4a, 60 HAP). Importantly, we found YFP-TOB1 co-localized with the tonoplast marker protein CFP-Vac-ck marker³⁷ (Fig. 4b) and the tonoplast marker dye FM4-64³⁸, surrounding the vacuole lumen marker dye BCECF³⁹ (Supplementary Fig. 5b), similar to the observations in roots²⁴. Therefore, we propose that tonoplastic TOB1 in MCs sequesters IBA into vacuoles and thus functions to terminate PIN2 accumulation to the apical PM of MCs.

**Fig. 4: TOB1-mediated vacuole internalization of IBA terminates with PIN2 polar accumulation at cotyledon’s marginal cells.**

Vacuolar IBA importer TOB1 functions to downregulate PIN2 levels in cotyledons

If TOB1 functions to terminate PIN2 accumulation, then its accumulation is expected to correlate with PIN2 accumulation/disappearance spatiotemporal patterns. Thus, we imaged cotyledons expressing translational reporters for both PIN2-GFP (PIN2p::PIN2-GFP) and YFP-TOB1 (TOB1p::YFP-TOB1) before, during and after PIN2 accumulation in MCs. YFP/GFP signal was accurately separated by means of spectral microscopy⁴⁰. We found TOB1 signal showed progressive tip-to-base accumulation along the cotyledon’s margin while PIN2 signal disappears in the same direction (Fig. 4c, white vs red arrows). Although we presumed this is part of a continuous process, we noted 3 discrete stages. First, YFP-TOB1 showed weak signal near the cotyledons tip while PIN2-GFP was highly accumulated along the entirety of the MCs file (Fig. 4c, 48 HAP). Second, at 60 HAP, YFP-TOB1 consolidated into a ring-like subcellular signal close to the cotyledon tip, while PIN2-GFP disappeared at the tip but remained at the base (Fig. 4c, 60 HAP). Last, PIN2-GFP was depleted from MCs, whereas YFP-TOB1 was strongly accumulated, opposite to what was found 24 h earlier (Fig. 4c, 72 HAP). These results indicate that PIN2 and TOB1 accumulate in MCs in an opposing manner.

Next, we tested if TOB1 is functionally required to downregulate PIN2 by introgressing PIN2-GFP into two loss-of-function alleles, tob1-1 and tob1-3 (Supplementary Fig. 7a). Both point (tob1-1) and insertional mutation (tob1-3) increased cotyledon surface areas (Supplementary Fig. 6a). Additionally, both tob1 mutants showed more expanded epidermal cells when compared to equivalent wild-type cotyledons (Supplementary Fig. 6b–d). As expected, absence of TOB1 causes a 25-30% increase in PIN2 levels in 60 HAP cotyledons (Fig. 5a) and also a delayed PIN2 downregulation observed at 72 HAP (Supplementary Fig. 6e). To test the biological significance of this increment on PIN2 levels, we evaluated the auxin response distribution pattern as reflected with DR5::GUS expression. In both tob1 alleles, DR5::GUS signal was enhanced at the tip and at the edges compared to Col-0 wild type (Fig. 5c). Increased DR5::GUS signal in tob1 was indicative of increased upwards auxin transport and reflecting the inability to prevent IBA to IAA conversion by IBA vacuole storage (red dashed lines and arrowheads in Fig. 5c). Interestingly, in root tips the PIN2-GFP levels remain unaltered in tob1-1 and tob1-3 mutants (Supplementary Fig. 7b, c). Additionally, we did not find evidence for genetic interaction between TOB1 and PIN2 for controlling root gravitropism, as tob1-1 eir1-1 double mutant shows a similar agravitropic response as eir1-1 mutant (Supplementary Fig. 7d–e). Consistent with these findings, we found that TOB1 and PIN2 are not accumulated in the same cell type in roots (Supplementary Fig. 7f), further corroborating the notion that the TOB1 regulation of PIN2 levels is a cell-autonomous phenomenon. Thus, our results showed that in cotyledons but not in roots TOB1 functions to downregulate PIN2 levels most likely by sequestering IBA into vacuoles and consequently reducing cytoplasmic IAA levels.

**Fig. 5: TOB1 regulates PIN2 levels in Arabidopsis cotyledon marginal cells.**

Auxin feedback regulates TOB1 expression

The accumulation of TOB1 in the tonoplast (60 HAP) occurs after PIN2-mediated auxin flow throughout the MCs has started (24-36 HAP). Thus, we hypothesized that once auxin accumulates to relatively high levels it regulates TOB1 expression to terminate PIN2-mediated auxin transport. To test this, we treated cotyledons with incipient TOB1 levels (at 24 HAP) with 100 nM IAA for 4 h. Exogenously applied auxin resulted in a ~ 30% increment in TOB1 levels (Fig. 5e, f). Under auxin treatments, YFP-TOB1 levels increased along the entire margin (white arrowhead Fig. 5e) but were much higher in those cells closer to the tip (red arrows). This finding is consistent with the above-described tip-to-base wave in the expression of YFP-TOB1 (Fig. 4d, red arrowheads). In silico analysis of the TOB1 promoter revealed several auxin-related motifs, listing ARF7, ARF8, and mARF34 (ARF7/ARF19 paralog in maize) with high confidence in two different algorithms (Supplementary Fig. 7a). These observations suggest auxin may induce TOB1 expression directly, in addition to cytokinin, a well-known TOB1 regulator²⁴. Alternatively, auxin may regulate TOB1 gene expression indirectly through auxin-cytokinin interaction. Altogether, we propose that PIN2-transported auxin self-terminates PIN2 expression and auxin transport toward the cotyledon tip by activating the accumulation of TOB1, which further sequesters IBA into vacuoles.

Discussion

In general, the developmental program of an organism encompasses a continuum of discrete phases, and therefore relies on transient cell-surface signals, transient intracellular signaling structures^41,42 and transient cellular arrangements⁴³. Here we show that a transient self-organizing PIN2-based auxin flow takes place in the cotyledon’s margin cells, contributing to the cotyledon’s tip auxin maxima and the PC interdigitation. In leaf, PIN1, PIN3, PIN4, and PIN7 function in vasculature development independently from PIN2⁴⁴. Future research needs to demonstrate whether PIN2 establishes tip auxin maxima independently or collaboratively with other PINs.

Self-organizing transient signals, similar to the one described here, may be instrumental during different developmental transitions in plants and animals. Besides leaf development, auxin-related self-organizing transient signals may be involved in phyllotaxis⁴⁵, root elongation, and root branching by regulating the cell proliferation to differentiation transition⁴⁶, or in the transition to rapid hypocotyl elongation during germination⁴⁷. In animals, self-organized transient signals might control the circumferential actin filament bundles during the embryo to elongated worm transition in C. elegans⁴⁸ or the formation of the multicellular rosettes in the epithelial-mesenchymal transition (EMT) during mammalian embryo development⁴⁹ or mesenchymal invasion during cancer metastasis⁵⁰.

Remarkably, this transient auxin flow is generated and dismantled within a 48-hour time frame by positive and negative feedback loops, respectively. First, the transient auxin flow is self-promoted by turning-on PIN2 accumulation through the biosynthesis of IBA-derived auxin. Subsequently, the local high auxin at the tip generated by the auxin flow induces TOB1 expression, which import IBA into vacuoles, terminating PIN2 accumulation (Fig. 6). This self-organizing mechanism secures a pulse of auxin flow towards the cotyledon tip, and thus our findings uncover an exciting mechanism for the generation of transient developmental signals.

**Fig. 6: A working model for self-organizing transient auxin flow at cotyledon’s margins.**

Our findings also raise some interesting questions about auxin synthesis, auxin flows, and signaling during developmental transitions. First, what is the advantage of employing IBA over other IAA sources? We showed the IBA-to-IAA pathway has a predominant role in MCs and gene expression indicates a preference for this pathway in the entire matured embryo. Early development and expansion of the matured embryo is fueled through fatty acid β-oxidation, a process similar to IBA-to-IAA conversion⁵¹. Thus, the pathway for auxin synthesis may be tightly linked to the seedling energy sources. Future studies could be directed at determining if and how auxin synthesis pathway preferences switch once seedlings become autotrophic. Second, what is the initial signal that activates the IBA-dependent auxin biosynthesis? The signal must be intrinsic as it is produced during seed imbibition. It could be a mechanical signal resulting from increasing turgor pressure and/or initial cotyledon expansion. Third, what is the signal establishing apical PIN2 polarity along MCs?. Different options can be considered; a pre-established tissue-wide polarity field^52,53, a combination of gene expression waves^54,55 and positive auxin feedback^56,57, or a nested self-organizing system between localization and auxin flow, as subjected by PIN1 during PC morphogenesis^4,58,59. In fact, PINs are transiently auxin-induced⁶⁰ and present during leaf serration^53,61.

Leaf borders, in particular leaf serration, have been found critical for the evolution of leaf shape diversity^53,61. As we have observed polar PIN2 in cotyledons, it would be interesting to decipher if the PIN2-based auxin transient flow and its contribution to PC morphogenesis could work as a link between PC shape diversity⁶² and leaf shape diversity^63,64.

Finally, it is well known that roots and stems show coordinated localization of polar PINs but usually in a multi cell-type context and without a clear start of the process. In contrast, cotyledon PIN2 localization occurs in only one cell type, the marginal cells, and the whole process is activated specifically at 24 HAP, moving auxin tip-wards throughout 10-15 cells. Thus, we propose that transient auxin flow at MCs might bring an exceptional opportunity to study the mechanisms underlying PIN-based auxin flow.

Methods

Plant growth conditions

All plants were grown at 22 °C under a 16 h light/ 8 h dark photoperiod with 85-90 µmol/s^-2 of light intensity. In vitro growth was carried out in 0.5X MS basal salt medium with vitamins, 1% sucrose, pH 5.7–5.8 and 0.8% Agar (Sigma-Aldrich® A1296). Hours After Plating (HAP) are counted from the moment the seeds are transferred to the growth conditions after stratification.

Plant materials

Arabidopsis Columbia (Col-0) ecotype is the genetic background for the subsequent lines, including mutants eir1-1/pin2/CS8058³², pin2-T/SALK_122916C³³, tob1-3/SALK_205450²⁴, tob1-1²⁴, as well as the reporter lines TOB1p::YFP-TOB1²⁴, PIN2p::PIN2-GFP³¹, TAA1p::GFP-TAA1^11,36, 35S::GFP-LTI6b⁶⁵, and CFP-Vac-ck/CS16256³⁷. R2D2 is in the Columbia-Utrecht ecotype³⁴. Dual reporters and reporters in mutant backgrounds were generated by introgression. Upon crossing reporters with a mutant background, homozygous mutations were confirmed in F2, and non-segregating lines for the reporter were obtained by screening several F3 lines. Only non-segregant F3 lines were used throughout the study. Primer sequences and genotyping strategy can be found in Supplementary Table 1.

Seed sterilization

For experiments analyzing specific developmental time points, seeds were bleach-sterilized as follows: 10 min agitation with 30% bleach in water with 0.1% TX-100 (Triton X-100), followed by rinsing 5 times with sterile water and subsequent stratification for 48 h at 4 °C in dark. Seed sterilization is crucial for the reproducibility of the developmental timepoints described in this manuscript. For routine experiments, sterilization was performed overnight using the vapor-phase method (Clough and Bent, 1998) carried out by simply placing seeds in a desiccator next to a beaker containing 80 mL of bleach, with 1 mL of concentrated HCl carefully added into the bleach⁶⁶.

Developmental stages

Using the bleach sterilization protocol and our described plant growing conditions, we observed the following hallmarks in seedling growth: at 0 HAP, a swollen seed coat; at 24 HAP, a longitudinal rupture in the seed coat; at 36 HAP, radicle protrusion but cotyledons remain inside the seed coat; at 48 HAP, cotyledons were greenish and some were covered with their seed coat and root hair-like structures became noticeable at the root-hypocotyl junction; at 60 HAP, green ellipsoidal cotyledons were rarely covered by the seed coat and form a ~ 90° angle between them; at 72 HAP, green globular-shaped cotyledons were flat with a ~ 180° angle between them. Hours After Plating (HAP) time is counted from the moment seeds are placed in growing conditions after stratification. Developmental time in ech2ibr10 mutants was timed as hours after root protrusion, as these mutants showed intrinsic delayed germination.

Vector construction

The ECH2 upstream regulatory region was amplified using promECH2-EcoRI, (GAATTCGGTTAGATATAGAAATGACGG) and promECH2-PmlI (CACGTGCCGATCAGGATTAGAGCTCAGC) primers. The resulting 577-bp product, spanning the region between ECH2 and its most upstream neighboring gene, was then cloned into the pCR4 vector (Life Technologies) to create pCR4-ECH2prom. The ECH2 upstream regulatory region was excised from pCR4-ECH2prom using EcoRI and PmlI enzymes and subcloned into the pMCS:YFP-GW⁶⁷ vector to create pECH2:YFP-GW. The ECH2 cDNA was recombined from pENTR-ECH2³⁰ into the pECH2:YFP-GW vector using LR Clonase (Life Technologies) to form pECH2::YFP-ECH2, which expresses an N-terminal YFP fusion with ECH2 driven by the ECH2 upstream regulatory region.

Imaging and microscopy

Adaxial sides of dissected cotyledons were analyzed. In the early stages of cotyledon development, careful removal of seed coat was necessary to dissect the cotyledon. All images were acquired using laser scanning confocal microscopes (20X objective, NA = 0.75–0.8). Specifically, ablation experiments were performed using either a Leica TCS SP5 or a Leica Stellaris 8 microscope. Linear unmixing was conducted using a Leica TCS SP8 X DLS or a Zeiss 880 microscope. Fluorescent protein (FP) signals were captured using the following settings: propidium iodide (PI; excitation 543/561 nm, emission 580–660 nm), GFP (excitation 488 nm, emission 505–535 nm), and YFP (excitation 488/514 nm, emission 492-530 nm or 520–570 nm, respectively). PC shape analysis was performed with PaCeQuant software as described before^6,68.

Linear unmixing

Signal separation of GFP/AF (autofluorescence), GFP/YFP and GFP/PI (propidium iodide) was performed by linear unmixing of spectral images⁴⁰ acquired with either the Leica TCS SP8 X DLS, Zeiss LSM 880 Elyra or Zeiss LSM 980 systems, generating identical results. In all cases, reference emission spectra are obtained from samples without mixed signals. For example, in the unmixing of GFP/AF (autofluorescence), a spectrum is obtained from the tip of wild type cotyledons (autofluorescence), while a GFP spectra is obtained from PIN2-GFP signal in the cotyledon’s margins of a stable transgenic line.

For the Leica TCS SP8 X DLS, signal separation was performed using lambda mode with a 20X/NA 0.75 objective, excitation with a 488 nm white pulse laser scanning from 490 nm to 630 nm, and detection with HyD detectors. The images were processed using the linear unmixing algorithm in LASX software. For the Zeiss 880, GFP/YFP and GFP/PI signal separation was performed using spectral imaging mode with a 20X/NA 0.8 objective, MBS 488, excitation with a 488 nm argon-ion laser and a 32-channel GaAsP spectral detector capable of simultaneous detection of 8.9 nm bands of signal ranging from 410 nm to 695 nm. For GFP/AF unmixing a Zeiss LSM 980 with 25X/NA 0.8 objective was used in lambda mode, excitation with 488 nm diode laser, and sequential detection with a 4-channel GaAsP spectral detector. For GFP/YFP and GFP/PI separation, we used a detection window of the 16 channels between 490–632, whereas for GFP/AF unmixing, a detection window of 15 channels between 491 nm and 623 was used. The images were processed using the linear unmixing algorithm in Zeiss ZEN software.

Image analysis

Intensity measurements and circularity [4π ·(area/perimeter²)] were obtained using Image J 1.40 (Wayne Rasband, NIH, USA) within a region of interest (ROI) as indicated in each figure legend. Circularity is an isoperimetric quotient, which compares the area of a shape, in our case a pavement cell, against the area of a circle with the same perimeter. Higher circularity values represent less lobing, as they are closer to a circle shape⁶⁹. To observe direct correlation with lobing, we defined interdigitation as 1-circularity. For cell border detection, dissected embryos were incubated either in 100 µg/mL PI for 10 minutes at room temperature (older cotyledons) or for 10 minutes at 4°C with 5 µM FM4-64 (younger cotyledons) prior to imaging. For apical PIN2-GFP abundance, we quantified the fluorescence at the apical region and normalized it against the background signal. For PIN2-GFP protein polarity, we quantified the fluorescence intensity at the apical side relative to a region of similar area at the middle of the lateral side for each MC. Similar results were obtained regardless of whether the lateral side was located closer to the tip or the base. The fluorescence intensity was obtained in Fiji as the integrated density from z-stack maximum projection confocal images. Both apical and lateral intensities were normalized against the average intensity of 3 randomly selected ROIs in the background.

GUS (beta-glucuronidase) activity assays

GUS staining assays using X-gluc as a substrate, also known as histological assays, were performed with slight modifications based on previous reports^70,71. In brief, plant samples were fixed in cold 80% acetone for at least 2 h, which could be extended overnight. Subsequently, they were vacuum infiltrated with the GUS solution for 10 min and incubated in the dark at 37˚C for 0.5 h. The GUS solution was prepared as described previously⁷⁰. Alternatively, 2-4 h incubation was necessary without vacuum infiltration. After staining, cotyledons were “cleared” in 70% ethanol with agitation overnight to remove the chlorophyll. Finally, cotyledons were dissected and observed under a light microscope.

GUS activity assays in intact plant tissue with 4-MUG as a substrate, also known as fluorometric assay, were performed with adaptations from previous reports^72,73. Samples (a pair of cotyledons) were incubated in 96-well microplates (one sample per well) containing 150 μL lysis buffer (50 mM sodium phosphate, pH 7.0, 10 mM EDTA, 0.1% Triton X-100). As suggested in the referenced literature, the entirety of the sample tissue must be completely submerged in the buffer for substrate penetration. Subsequently, 1 mM 4-MUG was added to initiate the reaction, which was then incubated at 37˚C for 90 min before being terminated with 50 μL of 0.2 M Na₂CO₃. The substrate 4-MUG is converted to the fluorescence product, 4-MU. To measure 4-MU fluorescence intensity, a multi-mode plate reader SpectraMax iD5 was used under the following specific wavelengths; 365 nm for excitation and 455 nm for emission. A standard curve generated with 4-MU (in lysis buffer plus stop solution) corroborated linearity (R² = 0.99) for this assay under our experimental conditions within the range of 10 nM to 1000 nM of 4-MU. GUS activity is expressed as nanomolar units of 4-MU per sample.

Chemical treatments

Exogenous treatments with IAA (100 nM, EtOH) were performed by directly diluting them in liquid growth media without phytoagar. To inhibit the IPyA auxin-biosynthetic pathway, a two-chemical cocktail containing 50 µM Yucasin and 10 µM L-Kyneurin was used to ensure maximum inhibitory activity⁷⁴. Stock solutions for Yucasin (10 mM, DMSO) and L-Kyneurin (50 mM, DMSO) were directly diluted in the growth medium before plating. At 24 HAP, seedlings were transferred and incubated for 12 h. in the presence of the chemicals. For vacuole markers, seedlings at 60 HAP were treated with 5 µM FM4-64 for 10 min and 10 µM BCECF-AM for 1 h in darkness. After incubation samples were washed for 30 sec before mounting for observation. FM4-64 (Stock of 10 mM, DMSO) and BCECF (Stock of 10 mM, DMSO) were directly diluted in liquid growth medium without agar.

Laser ablation

Seedlings of 35S::GFP-LTI6b⁶⁵ at 48 HAP were incubated with 100 µg/mL with Propidium Iodide (PI) in a microscope slide. A UV 405 nm pulsed laser (300 µW power) at 100% laser intensity was used for cell ablation in a Leica SP5 confocal system or Leica Stellaris 8. For LeicaSP5, each cell was irradiated for 5 sec in maximum optical zoom. PI penetrating the cell borders and marking the nuclei demarcated successful ablation. The whole file of marginal cells was ablated within 2-3 minutes. For Stellaris 8, a small window at maximum optical zoom was assigned to each MCs and a sequential irradiation was initiated using the multi-acquisition mode. After ablation, seedlings were re-incubated in vitro in the same growing conditions for further analysis after 24 hours. Vasculature patterns were observed by mounting the mock and ablated samples in Image-IT™ Plant Tissue Clearing Reagent (Invitrogen, V11328).

In silico analysis

The sequence spanning from +1346 bp until transcription start site (TSS) of the TOB1 gene (AT1G72140) was analyzed using promoter analysis tool PlantRegMap. Additionally, the sequence extending +1000 bp upstream the TSS was examined using PSCAN, http://159.149.160.88/pscan, a widely used promoter analysis tool. The PINs and TAA/YUCCA gene expression analysis was obtained from published data^75,76 by submitting gene codes to Arabidopsis ePlant browser http://bar.utoronto.ca/eplant. Cell-type specific expression analysis data for PIN2, TOB1 was obtained from translatome data at http://efp.ucr.edu.

Statistical methods

All statistical analyses were performed using GraphPad Prism®. An unpaired two-samples two-tailed t-test, or a two-way ANOVA with Šídák’s test for multiple comparisons was used throughout the study, with a 95% confidence interval. Equal variance was assumed for the t-test and assessed with a post F-test. Unequal variances were subject to a Welch’s t-test or a Welch’s ANOVA with a Dunnett’s T3 multiple comparisons test. Statistical significance was defined as follows: ns (no significance) p-value > 0.05, * p-value < 0.05, ** p-value < 0.01, *** p-value < 0.001, **** p-value < 0.0001.

Reporting summary

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

Data availability

All data underlying this study are available from the corresponding author upon request. Source data are provided with this paper for the main and the Supplementary Figs. Gene accession numbers involved in this study: PIN2 = AT5G57090 and TOB1 = AT1G72140. Source data are provided with this paper.

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Acknowledgements

We acknowledge David Carter at UCR Microscopy Core for technical assistance with Zeiss 880 microscopy, Lei Shi at FAFU-UCR center for technical assistance with Leica SP8 microscopy, and Olivier Brun for technical assistance with Leica Stellaris 8. We are thankful to Dr. Yi Tao for providing published material, TAA1p::GFP-TAA1. This work was in part supported by the National Institute of General Medical Sciences to Z.Y. (GM081451), the National Science Foundation to X.C. and Z.Y. (1853698) and the National Natural Science Foundation of China Grants to Z.Y (3241101698).

Author information

Patricio Pérez-Henríquez & Jaimie Van Norman
Present address: Department of Molecular, Cell and Developmental Biology, University of California, Los Angeles, CA, 90095, USA
Marta Michniewicz
Present address: Bayer Crop Science, 700 W Chesterfield Pkwy W, Chesterfield, MO, 63017, USA

Authors and Affiliations

Institute of Integrated Genome Biology, and Department of Botany and Plant Sciences, University of California, Riverside, CA, 92521, USA
Patricio Pérez-Henríquez, Xue Pan, Carolyn Rasmussen, Jaimie Van Norman & Zhenbiao Yang
Haixia Institute of Science and Technology, Fujian Agriculture and Forestry University, Fuzhou, Fujian, China
Patricio Pérez-Henríquez, Shingo Nagawa, Wenxin Tang & Zhenbiao Yang
Faculty of Synthetic Biology, Shenzhen University of Advanced Technology, Shenzhen, Guangdong, China
Zhongchi Liu & Zhenbiao Yang
The Key Laboratory of Quantitative Synthetic Biology, Shenzhen Institute of Advanced Technology, Chinese Academy of Sciences, Shenzhen, Guangdong, China
Zhongchi Liu & Zhenbiao Yang
Department of Biological Sciences, University of Toronto-Scarborough, Toronto, ON, M1C1A4, Canada
Xue Pan
Department of Biology, Washington University, St. Louis, MO, 63130, USA
Marta Michniewicz & Lucia Strader
Department of Statistics, University of California, Riverside, CA, 92521, USA
Xinping Cui
Department of Biology, Duke University, Durham, NC, 27708, USA
Lucia Strader
Institute of Emerging Agricultural Technology, Shenzhen University of Advanced Technology, Shenzhen, Guangdong, China
Zhenbiao Yang

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Contributions

P.P.-H. carried out most of the experiments; Z.Y. and P.P-H. interpreted the results and designed experiments. M.M. cloned constructs for TOB1 and ECH2 supervised by L.S. X.C. helped with statistical analysis. J.V.N. co-supervised P.P-H. during the revision of this manuscript. S.N, W.T, X.P, Z.L, C.R., and J.V.N. contributed critical skills in microscopy. P.P-H. and Z.Y. wrote the manuscript. All authors commented on the manuscript.

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Pérez-Henríquez, P., Nagawa, S., Liu, Z. et al. PIN2-mediated self-organizing transient auxin flow contributes to auxin maxima at the tip of Arabidopsis cotyledons. Nat Commun 16, 1380 (2025). https://doi.org/10.1038/s41467-024-55480-8

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Received: 14 December 2022
Accepted: 09 December 2024
Published: 05 February 2025
DOI: https://doi.org/10.1038/s41467-024-55480-8