JULGI coordinates vascular development and leaf patterning through non-cell-autonomous regulation of miR165/166

Domain: biology
Published: 2026-01-10
Editorial outcome: major_revision

Abstract
The interplay between vascular development and leaf development is central to the evolutionary success of vascular plants, yet the underlying mechanisms remain poorly understood. Here, we identify the RNA-binding protein JULGI 1 (JUL1) as a key integrator of vascular and leaf development in Arabidopsis. JUL1 is transcribed in leaf veins, while the protein distributes throughout the leaf, where it binds G-rich regions of pri-MIR165a transcripts on the abaxial side. This interaction disrupts hairpin formation and inhibits DICER-LIKE 1 (DCL1)-mediated processing, reducing miR165/166 levels. Consequently, transcript levels of their targets, the Class III homeodomain leucine zipper (HD-ZIP III), increase, reinforcing adaxial-abaxial polarity. Our findings reveal a non-cell-autonomous mechanism by which vascular-derived JUL1 modulates miRNA biogenesis to coordinate leaf patterning with vascular development.

Introduction
The leaf is the primary photosynthetic organ of vascular plants, composed of distinct layers, including epidermis, mesophyll, and vasculature, each with unique developmental trajectories and physiological roles1–3. Coordinated interactions among these tissues are essential for establishing both the structural framework and functional capacity of the developing leaf. Early vascular plants like Rhyniophytes bore simple, leafless axes containing only vascular strands, and fossil evidence suggests that modern leaves evolved as a secondary innovation through lateral lamina expansion from these strands4. This evolutionary trajectory suggests that vascular tissues, beyond resource transport, may have functioned as structural and developmental axes, potentially contributing to leaf polarity and patterning. Developmental evidence supports this link: showing that the asymmetric distribution of specific molecular components establishes polarity, providing positional cues that guide pattern formation during organogenesis5. In early leaf development, primary adaxial–abaxial polarity is first established6–9. Based on positional information, PIN-FORMED (PIN) auxin efflux transporters direct auxin to accumulate along the central axis of the leaf primordium, where it induces the formation of the provasculature that is subsequently specified and differentiated10,11. As a result, the vasculature is formed with its xylem aligned toward the adaxial (upper) side and phloem toward the abaxial (lower) side, reflecting the underlying polarity12–14. Subsequently, surrounding tissues undergo coordinated spatial organization and differentiation aligning with the vascular axis and resulting in a pattern that closely mirrors the final geometry of the leaf lamina10,15. The frequent occurrence of defective leaf patterning following disruption of vascular patterning16,17, along with the extensive sharing of key regulatory factors between vascular and leaf development18–23, suggests their tight interconnection.

Inter-tissue communication is essential for structural organization during organogenesis and is often mediated by non-cell-autonomous regulatory mechanisms24. Such regulation requires the production of mobile signaling molecules, their spatially controlled movement between cells or tissues, and the modulation of gene expression in recipient cells. Non-coding RNAs (ncRNAs) have emerged as molecular mediators of this process25,26, functioning to regulate gene expression or sequester specific transcripts with high specificity and adaptability. In leaf development, abaxially expressed microRNA165/166 (miR165/166) maintains abaxial identity by restricting the activity of class III homeodomain leucine zipper (HD-ZIP III) transcription factors27–29. Conversely, adaxial miR390 triggers trans-acting short-interfering RNA–auxin response factor (tasiR-ARF) biogenesis from adaxially expressed trans-acting siRNA 3 (TAS3), leading to degradation of ARF3/4 transcripts and promoting adaxial growth30,31. Their opposing distributions are critical for vascular and leaf patterning. To ensure proper function in response to developmental and environmental cues, ncRNA expression and activity are tightly regulated by multiple protein classes, including transcription factors and RNA-binding proteins, which modulate ncRNA biogenesis, stability, and mobility32–34. However, the spatiotemporal mechanisms by which vascular development mediates underlying adaxial-abaxial polarity to shape leaf morphology, particularly through ncRNA regulation to integrate vascular and adjacent tissues during leaf formation, remain largely unknown.

Results
JULGI (JUL) proteins were previously shown to negatively regulate phloem development by binding to G-rich sequences within the 5′ untranslated regions (5′ UTRs) of the SUPPRESSOR OF MAX2 1-LIKE 4 (SMXL4) and SMXL5 transcripts, which encode positive regulators of phloem differentiation, and suppressing their translation35–37. This regulatory mechanism emerged with the evolution of core angiosperms, together with cis-acting G-rich elements that facilitate JUL binding35,38. These G-rich elements are predicted to form non-canonical secondary structures called RNA G-quadruplexes (RG4s). Notably, in Arabidopsis, in silico predictions have identified over 65,000 regions capable of forming RG4s39, suggesting that JUL1 may have additional roles beyond the translational suppression of SMXL4 and SMXL5 transcripts.

Besides their translational regulation, RNA-binding proteins can also regulate RNA processing40. Given that ncRNAs are not translated, any interaction between an RNA-binding protein and an ncRNA likely reflects a role in RNA processing. To explore a non-translational role for JUL, we looked for Arabidopsis ncRNAs with the putative minimal JUL1-binding sequence, KKGDGGGKW (with K meaning G or U; D meaning any ribonucleotide except C; and W meaning A or U). This sequence was previously identified from Systematic Evolution of Ligands by Exponential Enrichment (SELEX)-enriched sequences (Fig. 1a, top)35,41. We detected this sequence in 44 ncRNAs. We separately searched for ncRNAs with G-scores > 10, indicating high probability of RG4 formation42, returning a list of 446 ncRNAs. Of 24 ncRNAs overlapping between these two lists, we ultimately chose five pri-MIR165 and pri-MIR166 transcripts as potential JUL1 targets (Fig. 1a and Supplementary Data 1), given the evolutionary conservation of JUL proteins in core angiosperms and the JUL1-binding sequence in core angiosperms such as black cottonwood (Populus trichocarpa, a dicotyledonous tree), Arabidopsis (a dicot), and rice (Oryza sativa, a monocot) and its absence in the fern Selaginella moellendorffii and the moss Physcomitrium patens (Fig. 1b and Supplementary Data 2)35,38,43.

pri-MIRNAs are the primary transcripts from which mature 21–24 nucleotide (nt) miRNAs are produced in plants and animals44. Mature miR165/166 molecules regulate adaxial–abaxial patterning by cleaving transcripts encoding HD-ZIP III transcription factors, key regulators of adaxial-side formation in plants20,23,29,45,46. Arabidopsis has two MIR165 and seven MIR166 loci23,29, producing nine pri-MIR165 or pri-MIR166 transcripts, five of which possess JUL1-binding sequences (Supplementary Data 3). In these pri-MIR165 and pri-MIR166 transcripts, the JUL1-binding sequence and the G-rich elements lie 44 nt from the 5′ end, overlapping with the sequence reverse complementary to mature miR165/166 (Fig. 1c).

To investigate if JUL has additional roles beyond phloem differentiation, we closely examined the morphological changes in jul mutants. Notably, we observed markedly altered leaf morphology in the knockout plants, with jul1 and jul1 jul2 mutants showing downward-curling, conical leaves (Supplementary Fig. 1a), a typical phenotype of increased abaxial leaf growth in Arabidopsis6,46–48. To quantify leaf curling, we measured the ratio between visible leaf area and flattened leaf area49. This ratio was significantly higher in jul1 and jul1 jul2 relative to the wild-type Col-0, and higher in jul1 jul2 than in jul1, suggesting a dosage-dependent effect (Supplementary Fig. 1b). In addition, the jul1 jul2 mutant showed higher expression of the abaxial marker genes FILAMENTOUS FLOWER (FIL) and KANADI (KAN) and lower expression of the adaxial marker genes PHABULOSA (PHB) and REVOLUTA (REV) compared to Col-0 (Supplementary Fig. 1c). These data indicate that JUL is involved in leaf adaxial–abaxial patterning.

To determine whether JUL affects the expression of MIR165/166, we measured the levels of pri-MIR165a, pri-MIR166a, and mature miR165/166 in Col-0, jul1, and jul1 jul2. We used pri-MIR166c and pri-MIR166d as internal negative controls that lack the putative minimal G-rich sequences in primary MIR165 and MIR166 transcripts, key regulators of adaxial–abaxial leaf polarity. In jul1 and jul1 jul2, pri-MIR165a and pri-MIR166a levels were lower than in Col-0, whereas those of pri-MIR166c and pri-MIR166d were comparable to Col-0 levels. Mature miR165/166 abundance gradually rose in the order Col-0, jul1 and jul1 jul2 (Fig. 1d). These results indicate that pri-MIR165 and pri-MIR166 are likely targets of JUL for the regulation of leaf adaxial–abaxial patterning. To further assess the functional connection between JUL and miR165/166 in leaf adaxial–abaxial patterning, we blocked miR165/166 activity using the Short Tandem Target Mimic (STTM) method50,51. Specifically, we expressed STTM165/166, a sequence complementary to miR165/166 but lacking the three central nucleotides, under the control of the meristem-specific RIBOSOMAL PROTEIN S5 (RPS5A) promoter in Col-0 and jul1 jul2 plants (Supplementary Fig. 2). In the shoot apices of jul1 jul2 proRPS5A:STTM165/166, miR165/166 levels were reduced by approximately 40% relative to jul1 jul2 (Fig. 1e). Whereas the leaves of jul1 jul2 plants were curled downward and conical-shaped, with a small rosette diameter, jul1 jul2 proRPS5A:STTM165/166 plants had flattened, round leaves, with a larger rosette diameter, resembling Col-0 plants (Fig. 1f). Additionally, the ratio of curled leaf area to flattened leaf area was lower in jul1 jul2 proRPS5A:STTM165/166 plants than in jul1 jul2, returning to Col-0 levels (Fig. 1g). Consistent with these phenotypic changes, the abaxial marker genes FIL and KAN were expressed at lower levels, whereas the adaxial marker genes PHB and REV were more highly expressed, returning to levels similar to Col-0 (Fig. 1h, i). These findings suggest that elevated miR165/166 activity underlies adaxial-abaxial imbalance of jul1 jul2, and that JUL modulates miR165/166 levels to maintain adaxial–abaxial patterning.

Vascular bundles in leaves are arranged along the adaxial–abaxial axis, placing xylem on the adaxial side and phloem on the abaxial side14. Similar to leaf curling, the more extensive phloem development observed in jul1 jul2 petioles and inflorescence stems relative to Col-0 returned to Col-0 levels in jul1 jul2 proRPS5A:STTM165/166 plants (Supplementary Fig. 3a–e). This morphological rescue was associated with lower expression of the phloem marker gene ALTERED PHLOEM DEVELOPMENT (APL) in jul1 jul2 proRPS5A:STTM165/166 than in jul1 jul2. Notably, the expression levels of the xylem marker gene XYLEM CYSTEINE PEPTIDASE 1 (XCP1) and the cambium marker gene TDIF RECEPTOR [TDR, also reported as PHLOEM INTERCALATED WITH XYLEM (PXY)] were not significantly different between jul1 jul2 and jul1 jul2 proRPS5A:STTM165/166 (Supplementary Fig. 3f). These results suggest that miR165/166 may contribute to JUL-mediated phloem development.

Phloem-derived JUL1 influences leaf adaxial–abaxial patterning via miR165/166
To investigate the spatial expression patterns of JUL1, we examined publicly available single-cell RNA-seq datasets from the Leaf SC Atlas52. JUL1 expression was predominantly enriched in cells engaged in the cell cycle, as well as in phloem-related cells, such as companion cells, sieve elements, and phloem parenchyma, where the phloem marker gene APL was also expressed. In contrast, JUL1 transcripts were not detected in abaxial pavement cells, adaxial pavement cells, or guard cells, in which the leaf patterning markers FIL, KAN, PHB, and PHV were expressed (Supplementary Fig. 4). To validate the spatial distribution of JUL1 and pri-MIR165a, we analysed their expression patterns in leaf primordia using green fluorescent protein (GFP) transcriptional fusion lines (proJUL1:3xGFP and proMIR165a:GUS-GFP) and a translational fusion line (jul1 jul2 proJUL1:JUL1-GFP). Consistent with previously published data28, we detected GFP fluorescence indicative of MIR165a transcriptional activity in the abaxial domain of young leaf primordia in proMIR165a:GUS-GFP transgenic seedlings (Fig. 2a, b and Supplementary Figs. 5a, e and 6a). By contrast, we observed JUL1 transcriptional activity specifically in the midvein of leaf primordia in proJUL1:3xGFP seedlings (Fig. 2a, b and Supplementary Figs. 5b, f and 6a). Notably, JUL1-GFP accumulated throughout the leaf primordium of jul1 jul2 proJUL1:JUL1-GFP seedlings (Fig. 2a, b and Supplementary Figs. 5c, g and 6a). These data suggest that JUL1 may function in a non-cell-autonomous manner, potentially influencing target gene expression beyond the transcriptional domain of its encoding gene.

To test the above notion, we examined whether tissue-specific expression of JUL1 could rescue the downward-curling, conical leaf phenotype of jul1 jul2 mutant plants. To this end, we placed JUL1 transcription under the control of promoters specific to the abaxial region (FIL), adaxial region (BLADE ON PETIOLE 1 [BOP1]), or vasculature (APL) and introduced each construct into the jul1 jul2 background (Fig. 2c, d). In all constructs, the JUL1 coding sequence was cloned in-frame and upstream of the sequence for a StrepII tag. In each case, ectopic JUL1 expression was sufficient to rescue the mutant leaf phenotype (Fig. 2c, e). Ectopic expression of JUL1 in the abaxial or vascular domain, driven by the FIL or APL promoter, respectively, fully restored the leaf phenotype to that seen in Col-0 plants; JUL1 expression in the adaxial domain driven by the BOP1 promoter also alleviated the curling phenotype typical of jul1 jul2, although to a lesser extent. Leaves of jul1 jul2 proBOP1:JUL1 plants exhibited more pronounced curling than those of Col-0, jul1 jul2 proFIL:JUL1, and jul1 jul2 proAPL:JUL1 plants. This morphological rescue was accompanied by changes in the expression levels of abaxial marker genes (FIL, KAN) and adaxial marker genes (PHB, REV). Compared to Col-0, jul1 jul2 plants expressed the abaxial marker genes to higher levels and the adaxial marker genes to lower levels. By contrast, these marker genes were expressed at levels comparable to Col-0 in the phenotype-rescued transgenic lines (Fig. 2f, g). Together, these results indicate that JUL1 functions as a non-cell-autonomous regulator of adaxial–abaxial patterning.

JUL1 specifically binds to the G-rich region of pri-MIR165a
To assess whether JUL1 directly binds to pri-MIR165/166, we conducted RNA electrophoretic mobility shift assays (REMSAs). As JUL binds to G-rich elements predicted to form RG4 structures35, we designed single-stranded RNA probes containing the JUL1-binding sequence (G-block#1) and two adjacent guanine-rich elements (G-block#2 and G-block#3). These sequences have the potential to form RG4 structures in vitro but do not match the canonical RG4-forming motif (5′-G≥3 N 1–7 G≥3 N 1–7 G≥3 N 1–7 G≥3 -3′), suggesting a limited propensity for stable RG4 formation (Supplementary Fig. 7). Using GST-tagged JUL1 and GST-JUL1(RA), a dominant-negative form of JUL1 lacking RNA-binding activity35, we tested binding to the G-rich elements of pri-MIR165a and pri-MIR166a, both of which are affected in julgi mutants (Fig. 1d). GST-JUL1 retarded the mobility of both G-rich elements, whereas GST-JUL1(RA) did not (Supplementary Fig. 8b), with JUL1 showing a stronger binding affinity to the G-rich element of pri-MIR165a than pri-MIR166a. Consistent with these results, bead-surface binding assays demonstrated condensation of Cy5-labeled RNA probes at the liquid–solid interface of JUL1-coated beads. Condensation occurred with both probes but was absent with JUL1(RA)-coated beads (Supplementary Fig. 8c). These results demonstrate that both pri-MIR165a and pri-MIR166a can directly interact with JUL1, but because pri-MIR165a exhibited a stronger binding affinity, we focused subsequent analyses on pri-MIR165a.

To further investigate whether JUL specifically binds to the three G-blocks within pri-MIR165a and assess the relative importance of each motif, we generated various G-to-A mutant probes (mutALL, mut#1, mut#2, and mut#3) in which all G nucleotides within each G-block were changed to A. Recombinant GST-JUL1 caused a mobility shift when incubated with the intact probe, confirming its binding activity, whereas recombinant GST-JUL1(RA) produced no shift (Fig. 3a). GST-JUL1 failed to shift the mutALL probe, in which all Gs in all three motifs were changed to As. The probes with individually mutated G-blocks (mut#1, mut#2, and mut#3) exhibited weaker shifts than the intact probe, with the strength of residual binding being mut#3, mut#2, and mut#1 from highest to lowest binding (Fig. 3a, b). To better pinpoint the binding site of JUL1 within pri-MIR165a, we conducted REMSA using radiolabelled probes and unlabelled (cold) competitors. Recombinant GST-JUL1 binding to intact radiolabelled probes was disturbed by increasing amounts of cold competitor probes. The intact probe showed the strongest competition capacity, whereas mutALL failed to disturb the protein–probe interaction when incubated with the intact probe or the full-length pri-MIR165a (Fig. 3c, d and Supplementary Fig. 8d, e). The competition capacity of the G-block mutants was highest for mut#3, followed by mut#2 and mut#1. These results demonstrate that JUL1 directly binds to three distinct G-blocks within the G-rich elements of pri-MIR165a. Although each G-block contributes differently to the binding affinity, all G-blocks are required for JUL1-binding.

To further examine the association between JUL1 and pri-MIR165a, we turned to an RNA monitoring system based on the hairpin from bacteriophage MS2 and MS2 coat protein in Arabidopsis protoplasts35,53. Accordingly, we appended 24 copies of the MS2 hairpin to full-length pri-MIR165a, enabling direct visualization of pri-MIR165a via binding of GFP-tagged MS2 coat protein in vivo. We co-transfected this RNA monitoring system into Arabidopsis protoplasts with a construct encoding mRFP-tagged JUL1 or mRFP-JUL1(RA). We observed co-localization of MS2-GFP with cytoplasmic compartments of mRFP-JUL1 when pri-MIR165a was fused with MS2 hairpins, but not with mRFP-JUL1(RA) (Fig. 3e, f), indicating that JUL1 associates with pri-MIR165a. Next, we performed an RNA immunoprecipitation (RIP) assay to directly assay the interaction between JUL1 and pri-MIR165a, using anti-HA antibodies on leaf extracts from Col-0 and two jul1 proJUL1:JUL1-HA complementation lines. Compared to Col-0, pri-MIR165a was highly enriched in the immunoprecipitates from jul1 proJUL1:JUL1-HA leaves, confirming the specific association of JUL1 with pri-MIR165a in vivo (Fig. 3g). Taken together, these results indicate that JUL1 directly binds to the G-rich region of pri-MIR165a in vitro and in planta.

JUL1 inhibits DCL1-mediated processing of pri-MIR165a
In the jul1 and jul1 jul2 mutants, pri-MIR165a levels were lower, whereas those of mature miR165/166 were higher (Fig. 1d), suggesting that JUL1 is involved in miRNA processing. During the processing of pri-MIRNA transcripts into mature miRNAs, DICER-LIKE 1 (DCL1) typically recognizes and cleaves hairpin-structured double-stranded pri-MIRNAs, producing a 21-bp miRNA/miRNA* duplex54,55. However, JUL1 is thought to bind single-stranded RNA through its RanBP2-type zinc finger (ZnF) domains35,56.

To explore whether JUL1 binding to pri-MIR165a prevents the formation of double-stranded (ds) RNA, we employed a bead surface binding system that captures a Cy3-labelled anti-probe onto streptavidin beads coated with a biotinylated, Cy5-labelled probe corresponding to the entire G-rich element with all three G-blocks. Beads coated with the probe were surrounded by red signal under confocal microscopy, and co-incubation with the anti-probe produced a green fluorescence signal in the absence of JUL1, indicative of dsRNA formation (Fig. 4a). Importantly, the Cy3 green signal intensity was much lower in the presence of JUL1, suggesting diminished co-localization of the probe and anti-probe or dsRNA formation. By contrast, fluorescence signals in the presence of JUL1(RA) were comparable to those in the absence of JUL1. Subsequently, we conducted Förster resonance energy transfer (FRET) analysis using the entire G-rich element (labelled with Cy5) and the anti-probe (labelled with Cy3) in the presence of JUL1 or JUL1(RA) (Fig. 4b). When we added the Cy5-labelled probe to the Cy3-labelled anti-probe, the FRET ratio rose in the absence of JUL1 and in the presence of JUL1(RA). However, the FRET ratio was significantly lower in the presence of JUL1 relative to the absence of JUL1 or the presence of JUL1(RA). These findings support the notion that JUL1 impedes hairpin-loop dsRNA formation by binding to single-stranded pri-MIR165a.

To investigate if JUL1 affects the interaction between pri-MIR165a and the dsRNA-binding domain (dsRBD) of DCL1, we conducted native RIP assays for pri-MIR165a using the dsRBD of DCL1 conjugated to an HA tag (dsRBD-HA) in the presence or absence of FLAG-tagged JUL1 (JUL1-FLAG). In protoplasts co-expressing JUL1-FLAG, pri-MIR165a levels co-immunoprecipitated with dsRBD-HA were significantly lower than those obtained in the absence of JUL1 (Fig. 4c). To confirm whether JUL1 impairs DCL1 activity toward pri-MIR165a, leading to its diminished processing into mature miR165, we conducted an in vitro pri-MIRNA processing assay. We used radiolabelled pri-MIR165a_WT and its mutant form pri-MIR165a_mut#3, to which JUL1 has a lower binding affinity, without altering the target specificity of mature miR165a. We performed the assay using nuclear extracts from Arabidopsis protoplasts overexpressing YFP-DCL1, along with increasing amounts of recombinant GST-JUL1 added to the extracts. Nuclear extract facilitated the production of a 21-nt mature miR165 from both the 199-nt pri-MIR165a_WT and pri-MIR165a_mut#3 (Fig. 4d). With more GST-JUL1 added, the abundance of processed miR165a from pri-MIR165a_WT diminished; however, pri-MIR165a_mut#3 was associated with a weaker suppression of mature miR165 production than pri-MIR165a_WT, at an identical concentration of GST-JUL1, indicating that JUL1 reduces the efficiency of DCL1-mediated processing of pri-MIR165a into mature miR165. Collectively, these results suggest that JUL1 interferes with the incorporation of pri-MIR165a into the dsRNA-binding domain of DCL1, possibly through competitive binding or steric hindrance, thereby suppressing miR165a biogenesis.

JUL1 regulates leaf adaxial–abaxial growth by suppressing miR165/166 levels
To establish the effects of JUL1 on miR165a biogenesis and its downstream targets, we co-transfected pri-MIR165a into Arabidopsis protoplasts with GFP reporters harbouring the GFP coding sequence cloned in-frame and downstream of the intact miR165/166-binding site of PHB transcripts (PHBBS-GFP) or a mutant in the miR165/166-binding site (mPHBBS-GFP)45,46 with JUL1 or JUL1(RA). In protoplasts expressing the PHBBS-GFP reporter, we observed JUL1-dependent increases in pri-MIR165a levels, PHBBS-GFP transcripts and GFP signals, concomitant with lower levels of mature miR165/166 (Fig. 5a, top). By contrast, in protoplasts expressing the mPHBBS-GFP reporter, GFP transcripts and GFP signals remained constant regardless of the amount of JUL1 added, whereas pri-MIR165a abundance rose and that of mature miR165/166 diminished in a JUL1-dependent manner. Unlike JUL1, JUL1(RA) did not affect mature miR165/166 levels, nor did it influence GFP transcript or GFP levels in protoplasts expressing either PHBBS-GFP or mPHBBS-GFP (Fig. 5a, bottom). To exclude the possibility that the observed phenotypic changes in jul1 and jul1 jul2 indirectly affect miR165/166 levels and to clarify the specific role of JUL1 in miR165/166 biogenesis, we assessed the abundance of pri-MIR165a, mature miR165/166, and the miR165/166 target transcripts PHB and REV in lines with JUL1-inducible expression in the jul1 background (jul1 LhGR-JUL1) and JUL1-inducible knockdown in the Col-0 background using an artificial microRNA (LhGR-amiR-JUL1). Compared to mock-treated seedlings, dexamethasone (DEX)-treated jul1 LhGR-JUL1 seedlings showed significantly higher levels of JUL1 transcripts and pri-MIR165a and lower levels of mature miR165/166 (Fig. 5b). Consistent with the lower miR165/166 levels, PHB and REV transcript levels were significantly higher in DEX-treated seedlings than in mock-treated seedlings. By contrast, DEX treatment of LhGR-amiR-JUL1 seedlings resulted in markedly lower levels of JUL1, PHB and REV transcripts and pri-MIR165a, alongside greater abundance of mature miR165/166 (Fig. 5c). These findings suggest that the curled-leaf phenotype and altered miR165/166 abundance in jul1 jul2 plants can be directly attributed to the regulatory role of JUL toward miRNA abundance, rather than secondary effects caused by greater phloem and abaxial development.

To further investigate the roles of JUL in miR165/166 biogenesis during leaf adaxial–abaxial patterning, we introduced the constructs proMIR165a:MIR165a_WT and proMIR165a:MIR165a_mut#3 individually into Col-0 and jul1 jul2. In the Col-0 background, proMIR165a:MIR165a_WT transgenic plants developed rosettes and leaves comparable to those of Col-0, with a similar degree of curling. By contrast, proMIR165a:MIR165a_mut#3 lines exhibited significant downward curling, indicating more pronounced abaxial growth (Fig. 5d, e). Notably, the curling phenotype in proMIR165a:MIR165a_mut#3 in Col-0 background closely resembled that of jul1 jul2. However, in the jul1 jul2 background, neither transgene further altered leaf morphology beyond the jul1 jul2 phenotype. To confirm that the curly leaf phenotype was associated with JUL-dependent miR165/166 biogenesis, we measured pri-MIR165a and miR165/166 levels. Both proMIR165a:MIR165a_WT and proMIR165a:MIR165a_mut#3 transgenes led to higher abundance of mature miR165/166 in all lines compared to their respective backgrounds (Supplementary Fig. 9). However, the ratio of mature miR165/166 to pri-MIR165a, an indicator of processing efficiency, was significantly higher in proMIR165a:MIR165a_mut#3 than in its wild-type Col-0, whereas the ratio in proMIR165a:MIR165a_WT was similar to that in Col-0 (Fig. 5f). By contrast, we observed no significant differences in this ratio in jul1 jul2 or its derived transgenic lines. These results indicate that JUL1-binding to pri-MIR165a is crucial for suppressing miR165/166 processing and that disruption of this interaction leads to enhanced miR165/166 biogenesis and thus to leaf curling.

Methods
Plant materials and growth conditions
Arabidopsis thaliana Col-0, jul1, and jul1 jul2 were used as wild-type and genetic backgrounds for the transgenic lines. The jul1 jul2 mutants were used as described38. The jul1 mutant was generated by backcrossing the jul1 jul2 with Col-0. All seeds were germinated and grown for 1 week in a media containing 1/2 Gamborg B5 salts (Duchefa), 1% sucrose, and 0.65% phytagel (Duchefa) (pH 5.6–5.7) under long-day conditions (16 h light/8 h dark) at 21 °C. Plants for phenotypic analyses and RNA extraction were transplanted to pots at 7 days after sowing (DAS) and grown under long-day conditions. Plants for protoplast isolation were grown under short-day conditions (10 h light/14 h dark) at 21 °C and fully expanded leaves of 4-week-old plants were used.

Plasmid construction for transgenic plants and protoplast transient expression assay
To generate the JUL1 complementation line, the JUL1 coding sequence (CDS) was cloned into the pCAMBIA1303 vector containing 3.7 kb of the JUL1 promoter and HA or GFP tags. To monitor the spatial expression of MIR165a and JUL1, we cloned 2.3 kb of the MIR165a promoter23 or 3.7 kb of the JUL1 promoter into the pCAMBIA1303 vector containing GUS-GFP or 3xGFP, respectively. To generate meristem specific miR165/166 suppression line, the STTM165/166 sequence51 was cloned into the pCAMBIA1303 vector containing 1.7 kb of the RPS5A promoter67. To generate tissue specific JUL1 expression lines, plasmid construction was performed using the Golden Gate cloning68. JUL1-StrepII, 3 kb of FIL promoter48, 3 kb of APL promoter69, and 3.7 kb of BOP1 promoter70 were separately PCR-amplified, and each PCR product was cloned to the universal Level 0 (Lv0) vector (pAGM9121). Lv0 vectors of JUL1-StrepII and various promoters were assembled to Lv1 vectors. Each Lv1 vector was assembled to an Lv2 binary vector (pAGM4723) with the antibiotic-resistant gene BAR containing an Lv1 vector (pICSL11017) and linker (pICH41744). To generate DEX-inducible JUL expression transgenic line, JUL-StrepII was cloned into pCR8/GW/TOPO vector. To generate DEX-inducible gene suppression transgenic line, artificial microRNA (amiR) targeting JUL was designed using WMD371. amiR precursor was generated by overlapping PCR using pRS300 vector as a template and cloned into pCR8/GW/TOPO vector. Recombination reaction between entry vector (pCR8/GW/TOPO) and destination vector (pB7WG6S LhGR)72 was carried out using Gateway® LR Clonase™ II enzyme mix (Invitrogen), yielding LhGR-JUL and LhGR-amiR-JUL constructs. All constructs were transformed into Agrobacterium tumefaciens GV3101, and the Arabidopsis plants were transformed using the Agrobacterium floral dipping method73.

Protoplast expression vectors containing HA, GFP, and FLAG tags were used as templates to construct plasmids for protoplast transient expression74. pri-MIR165a, 24X MS2 hairpin-fused pri-MIR165a, double stranded RNA binding domain of DCL1 (dsRBD)75 were amplified by PCR and cloned into the HA vector. PHBBS, mPHBBS were constructed by a primer annealing and cloned into the GFP vector. The JUL1 CDS was amplified by PCR and cloned into the FLAG vector. mRFP-JUL1, mRFP-JUL1(RA), MS2-GFP, JUL1-HA, and JUL1(RA)-HA constructs were previously described35. The primers for cloning are listed in Supplementary Table 1.

Transient expression in Arabidopsis protoplast
For MS2 monitoring system and GFP reporter assay, 4×104 protoplasts were transfected with 40 μg of total plasmid DNA composed of different combinations of the vectors and incubated for 6 h at room temperature. For the cytosolic condensation of JUL1 protein, the samples were treated with 0.01% NaN3 for 10 min. GFP and mRFP fluorescence were observed by a fluorescence microscope (LSM800, Zeiss). For RNA immunoprecipitation and pri-MIR165a processing assay, 8×105 protoplasts were transfected with 800 μg of total plasmid DNA for RNA immunoprecipitation and 400 μg of total plasmid DNA for pri-MIR165a processing, composed of different combinations of the vectors, respectively and incubated for 16 h at room temperature. RNA in GFP reporter assay was assessed by RT-PCR. RNAs in RNA immunoprecipitation and pri-MIR165a processing assay were measured by quantitative RT-PCR. Proteins in GFP reporter assay, RNA immunoprecipitation and pri-MIR165a processing assay were detected by horseradish peroxidase (HRP)-conjugated high-affinity anti-GFP (Santa Cruz Biotechnology) and anti-HA (Invitrogen) antibodies, respectively, and visualized using ECL solution (Pierce) and ImageQuant LAS 500 (Amersham) or X-ray film (AGFA) expose.

Recombinant protein purification
For protein purification of GST-fused recombinant proteins, GST-fused JUL1 and JUL1(RA) were expressed in Escherichia coli BL21 strain. Cells were grown in 500 ml of Luria broth (LB) medium at 37 °C, until they reached an optical density at 600 nm of 0.6 to 0.7 with shaking. After 1 mM isopropyl β-d-1-thiogalactopyranide (IPTG) treatment, cells were further incubated 20 h at 18 °C. The recombinant GST-fused proteins were purified by Glutathione Sepharose 4B (Cytiva) according to the manufacturer’s protocol. GST-fused JUL1 and JUL1(RA) vectors were previously described35.

RNA isolation and quantitative RT-PCR (qRT-PCR)
Total RNAs from shoots of 7-day-after-sowing (DAS) seedlings, leaves of 5-week-old plants, and mesophyll protoplasts of 4-week-old plants were isolated using TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. Reverse transcription of mRNAs and pri-MIR165/166 was carried out with 1 μg of total RNA, oligo(dT) primers, and ImProm-II reverse transcriptase (Promega) according to the manufacturer’s instructions. Reverse transcription of mature miR165/166 was carried out with 1 μg of total RNA, miR165/166-specific stem loop primer, and Superscript III reverse transcriptase (Invitrogen) by pulsed reverse transcription76. qRT-PCR was performed with CFX Connect Real-Time System (BIO-RAD) and the SYBR Premix ExTaq system (Takara) according to the manufacturer’s instructions. Primers for qRT-PCR are listed in Supplementary Table 1.

Circular dichroism (CD) assay
CD assay was adoted with minor modification from the previous study35. All CD spectra were obtained from a J-815 CD Spectrometer (Jasco) at 25 °C using quartz cuvettes with a 2.0 mm path length. Each spectrum was recorded over a wavelength range of 220 nm to 320 nm with a 50 nm min−1 scanning speed. The final spectrum encompassed the average of five scans of the same sample. Synthesized RNA oligonucleotides (2 μM) were slowly cooled from 95 °C to room temperature in a binding buffer [10 mM Tris-HCl (pH 8.0) and 1 mM MgCl2, either in the absence or presence of 100 mM KCl] to reach equilibrium prior to CD measurements. The re-natured RNA oligonucleotides containing the 100 mM KCl were combined with 1 μM JUL1 protein and incubated for 30 min before the CD measurement to observe the G-quadruplex structure of the RNA in the pri-MIR165a–JUL1 complex. To eliminate the influence of proteins and buffers, the spectra containing the JUL1 protein were corrected using 1 μM JUL1 in buffer as a baseline. The spectra of 1 μM JUL1 alone were also recorded to demonstrate that JUL1 has no significant effect on the spectrum of the RNA G-quadruplex.

RNA Electrophoretic mobility shift (REMSA) assay
RNA EMSA was adopted with minor modifications from the previous study35. For RNA EMSA with G-rich element, single-stranded RNA (ssRNA) oligonucleotides of the G-rich region of primary MIR165a (pri-MIR165a_G-rich), various G to A mutants (mutALL, mut#1, mut#2, and mut#3) and anti sensed counter part of G-rich element (anti-probe) were synthesized (Bioneer). For RNA EMSA of pri-MIR165a, full-length pri-MIR165a RNA was produced by in vitro transcription using DNA template, which was generated from PCR amplification of full-length pri-MIR165a containing the minimal T7 promoter. Transcription was carried out using the RiboMAX RNA Production System-T7 (Promega). Transcribed RNA was denatured and purified by gel electrophoresis using a 12% urea-TBE gel. RNA probes were labelled with [γ−32P]-ATP (10 mCi ml−1) by incubation with T4 polynucleotide kinase (New England Biolabs) for 30 min at 37 °C. The unlabeled radionucleotides were removed using an Illustra MicroSpin G25 column (Amersham). Radioisotope-labelled RNA probes were denatured by incubation at 95 °C for 5 min and rapidly cooled on ice. 0–40 μM of GST-JUL1, GST-JUL1(RA) proteins were incubated in a binding buffer [10 mM Tris-HCl (pH 8.0), 2.5% glycerol, 0.5 mM 1,4-dithiothreitol (DTT), 50 μg ml−1 BSA, 100 mM KCl, 250 μM EDTA, and 1 μg heparin] with 10 nM RNA probes for 20 minutes at room temperature. The reaction mixture was resolved on a 6–12% polyacrylamide gel in 0.5× TBE buffer. Gels were visualized on a Phosphor screen using a Typhoon FLA 9000 Phosphor Imager (GE Healthcare). RNA probes for RNA EMSA and primers for template amplification are listed in Supplementary Table 1.

Nuclear protein extraction
Sucrose gradient subcellular fractionation was performed as previously described with minor modification77. Briefly, YPF-DCL1 expressing Arabidopsis protoplasts were suspended in cell lysis buffer [20 mM Tris-Cl (pH 7.5), 250 mM Sucrose, 2.5 mM MgCl2, 25% glycerol, 20 mM KCl, 2 mM EDTA, 1% Triton X-100, 1 mM PMSF, 5 mM DTT, and protease inhibitor (PI; Roche)] and the nuclei is pelleted by centrifugation at 1500×g for 10 min at 4 °C. This pellet was gently washed 4 times with cell lysis buffer. Washed pellets were resuspended in nuclei resuspension buffer 1 [20 mM Tris-Cl (pH 7.5), 250 mM Sucrose, 10 mM MgCl2, 0.5% Triton X-100, 5 mM 2-mrcaptoethanol (BME), 1 mM PMSF, and PI]. The suspension was layered on the top of equal volume of nuclei resuspension buffer 2 [20 mM Tris-Cl (pH 7.5), 10 mM MgCl2, 1.7 M sucrose, 0.5% Triton X-100, 5 mM BME, 1 mM PMSF, and PI] and centrifuged at 16,000×g for 45 min at 4 °C. The supernatant was removed, and the pellet was lysed with nuclei lysis buffer [50 mM Tris-Cl (pH 7.5), 10 mM EDTA, 0.05% SDS, 0.1% Triton X-100, and PI] to obtain the nuclear fraction. The nuclear fraction was used for the in vitro pri-MIRNA processing assay.

In vitro pri-MIRNA processing assay
In vitro pri-MIRNA processing assay was performed as previously described with minor modifications75. Transcription was carried out with the DNA template of pri-MIR165a described above using the RiboMAX Large Scale RNA Production System-T7 (Promega) in the presence of [α−32P]-UTP (10 mCi ml−1), and the RNA probes were denatured and purified by gel electrophoresis using a 12% urea-TBE gel. 100 nM of pri-MIR165a RNA probes were incubated with an activity buffer [20 mM Tris-HCl (pH 7.0), 50 mM NaCl, 4 mM MgCl2, 5 mM ATP, and 1 mM GTP] and 2 U μl−1 RNasin RNase inhibitor (Promega) for 40 min at 37 °C, in the absence or presence of 2.5 μM of GST-DCL1 and 0 to 20 μM of GST-JUL1. After incubation, proteinase K (NEB) was added and incubated for 15 min. For gel electrophoresis, the samples were mixed with RNA Gel Loading Dye (Thermo Scientific), incubated at 95 °C for 5 min and cooled on ice. The incubated samples were separated on a 12% urea-TBE gel, fixed by fixation solution (20% ethanol and 5% acetic acid), and visualized on a Phosphor screen using a Typhoon FLA 9000 Phosphor Imager (GE Healthcare).

RNA immunoprecipitation
RNA immunoprecipitation was adopted with minor modifications from the previous study35. 1 mg of 4-week-old Col-0 and jul1 proJUL1:JUL1-HA leaves or 8×105 protoplasts were used for RNA immunoprecipitation sample. Cross-linking was done for 15 min with 1% formaldehyde under vacuum. After quenching with 0.125 M glycine for 10 min and brief rinse with 1x PBS (pH 7.4), leaves were frozen in liquid nitrogen and ground to fine powder. RNA-protein complexes in ground leaves were extracted using IP buffer [100 mM KCl, 2.5 mM MgCl2, 10 mM Tris-HCl (pH 8.0), 10% glycerol, 0.5% NP-40, 1 mM DTT, 100 U ml−1 RNasin RNase inhibitor (Promega), 25 μM MG132, and PI (Mini-complete tablet, Roche)]. After centrifugation at 13,000×g for 10 minutes at 4 °C, 300 μl of cell extracts were incubated with 1 μg of HA antibody (Roche) for overnight at 4 °C with gentle rotation. A 30 μl aliquot of the cell extracts was stored at −80 °C for later experiments. Dynabeads Protein G magnetic beads (Invitrogen) were washed three times in a washing buffer [100 mM KCl, 2.5 mM MgCl2, 10 mM Tris-HCl (pH 8.0), 10% glycerol, 0.5% NP-40, and 100 U ml−1 RNasin RNase inhibitor (Promega)], then the anti-HA-decorated extracts were incubated with 10 μl Protein G magnetic beads for 2 h at 4 °C with gentle rotation. The beads were then washed eight times with the washing buffer. Co-immunoprecipitated RNAs were isolated by TRIzol reagent (Invitrogen) and reverse transcription was carried out using SuperScript III (Invitrogen) according to the manufacturer’s instruction. Co-immunoprecipitated RNAs were analyzed by qRT-PCR and proteins were detected by horseradish peroxidase (HRP)-conjugated high-affinity anti-HA (Invitrogen) and anti-FLAG (Cell Signaling Technology) antibodies and visualized using ECL solution (Pierce) and ImageQuant LAS 500 (Amersham) or X-ray film (AGFA) expose. The primers for RNA immunoprecipitation are listed in Supplementary Table 1.

Bead surface binding assay
ssRNA oligonucleotides of the 5′ biotinylated and 3′ Cy5-labeled G-rich element (5′-[biotin]UUGAGGGGAAUGUUGUCUGGAU[Cy5]−3′) and 5′ Cy3-labeled anti-probe (5′-[Cy3]UCGGACCAGGCUUCAUCCCCCCCAA-3′) were synthesized (Bioneer). The RNA probes, with a concentration of 25 μM, were heated in a structure buffer [10 mM Tris-HCl (pH 8.0), 100 mM KCl, and 1 mM MgCl2] at 95 °C for 5 min and then gradually cooled to 25 °C. NeutrAvidin Agarose Resin beads (Invitrogen) were washed twice with the structure buffer and incubated with 2.5 μM of the G-rich element for 1 h at 4 °C. The RNA-bead complexes were washed twice with the structure buffer. In the absence or presence of 25 μM of GST-JUL or GST-JUL1(RA) proteins, 250 nM of anti-probe was added to RNA-bead complexes and incubated for 10 min at room temperature with occasional mixing. Cy3 and Cy5 fluorescence signals were observed with a confocal microscope (LSM800, Carl Zeiss). Cy3 and Cy5 were excited at the wavelengths of 488 nm and 640 nm, respectively. The emission wavelengths for Cy3 and Cy5 fluorescence were recorded at 400–640 nm and 650–700 nm, respectively. The RNA probes for bead binding assay are listed in Supplementary Table 1.

Förster resonance energy transfer (FRET) analysis
The ssRNA oligonucleotides in the bead surface binding assay were also utilized in the FRET analysis. The RNA probes, with a concentration of 25 μM, were heated in a structure buffer [10 mM Tris-HCl (pH 8.0), 100 mM KCl, and 1 mM MgCl2] at 95 °C for 5 min and then gradually cooled to 25 °C. The G-rich element was incubated overnight at 4 °C with GST-JUL1 or GST-JUL1(RA). The anti-probe was then added to RNA-protein mixture and incubated for 30 minutes at room temperature with occasional mixing. The final concentrations of each RNA probes and proteins were set to 2.5 μM and 25 μM, respectively. Fluorescence intensities of Cy3 and Cy5 were measured using a SYNERGY H1 multi-mode microplate reader (BioTek). The sample was excited at 520 nm, and the fluorescence emission intensity at 660 nm divided by the emission intensity at 560 nm was presented as a FRET ratio78. The RNA probes for FRET analysis are listed in Supplementary Table 1.

Adaxial–abaxial phenotypic analysis
The stem cross-section was performed as previously reported79. For stem cross section, 0.5-cm long sections of the inflorescence stem were harvested from the base when stem length was 20 cm and embedded in Spurr’s resin (Sigma-Aldrich), following the manufacturer’s instructions. Embedded stem samples were sectioned into 0.2 μm thickness using sliding microtome (RM2265, Leica). Cross-section images of stem samples were acquired using a light microscopy (Axioplan2, Carl Zeiss). Relative areas of the central and peripheral regions were calculated as the ratio of each region’s area to the total stem area. Transverse section of the mid-vein in leaf blades and petioles followed procedures as described previously80. Seventh leaves harvested at 21 DAS were embedded in Technovit 7100 resin (Kulzer Technik), following the manufacturer’s instructions. Embedded leaf samples were sectioned into 4 μm thickness using a rotary microtome (Microtome HM325, Thermo Scientific). Transverse-section images of leaves were acquired using a light microscopy (Carl Zeiss). Relative areas of xylem and phloem in mid-vein and petiole were calculated as the ratio of the area of xylem or phloem to the sum of xylem and phloem areas. Leaf and rosette phenotypes were observed at 28 DAS. To assess the extent of leaf curling, the visible curled areas and the flattened leaf areas were measured as described previously49. The visible curled leaf areas were estimated from photographs taken directly above the specimens. For the flattened leaf areas, same leaves are flattened under clear adhesive tape and photographed. ImageJ81 was used to calculate relative areas of central, peripheral, phloem, xylem, curled leaf, and flattened leaf region.

GUS staining
Seedlings at 7, 14, and 21 DAS were first cleared with 90% acetone for 1 h at 4 °C. Following a rinse with a phosphate buffer (pH 7.2), the cleared tissues were incubated overnight with GUS substrate solution [50 mM phosphate buffer (pH 7.0), 2.5 mM potassium ferrocyanide, 2.5 mM potassium ferricyanide, 0.1% Triton-X, 10 mM EDTA, 1 mg/ml 5-bromo-4-chloro-3-indolyl-β-d-glucuronidase (X-Gluc)] at 37 °C. Subsequently, remained chlorophyll in GUS-stained seedlings was removed with 70% ethanol.

RG4 analysis in Arabidopsis protein coding RNA, pre-MIRNA, lncRNA, and tRNA
Arabidopsis pre-MIRNA, lncRNA, tRNA, and protein coding RNA sequences were retrieved from miRbase (https://www.mirbase.org/)43, PLncDB (https://www.tobaccodb.org/plncdb/)82, tRex (http://combio.pl/trex)83, and TAIR10 (https://www.arabidopsis.org/), respectively. Probabilities of G-quadruplex formation (G-score) of Arabidopsis coding and non-coding RNA sequences were calculated from QGRS Mapper84. To analyze the putative minimal JUL1-binding sequence, RNA sequences were enriched by systematic evolution of ligands by exponential enrichment (SELEX)35. After enrichment, RNA sequences were analyzed by Multiple Em for Motif Elicitation (MEME; version 5.4.1)41 with following parameters [-rna -nostatus -mod zoops -nmotifs 5 -minw 6 -maxw 9 -objfun classic -markov_order 0].

Multiple alignments of pre-MIRNA 165/166 family sequences across plant species
Pre-MIRNA 165/166 family sequences of Populus trichocarpa, Arabidopsis thaliana, Oryza sativa, Selaginella moellendorffii, and Physcomitrium patens were retrieved from miRbase43. Multiple alignments of pre-MIRNA 165/166 family sequences were conducted using CLUSTAL W with default options in MEGA1185 and visualized using Unipro UGENE86.

Stem-loop structure prediction of pri-MIR165a
Arabidopsis pri-MIR165a sequence was retrieved from miRex 2.0 (http://www.combio.pl/mirex2)87 and stem-loop structure of pri-MIR165a was predicted using RNAfold web server (http://rna.tbi.univie.ac.at//cgi-bin/RNAWebSuite/RNAfold.cgi)88 with default options.

Statistical analyses
Statistical analyses of data were conducted using GraphPad Prism 9.0.

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

Data availability
Data are available within the Article, Supplementary Information or Source Data file. Source data are provided with this manuscript. Source data are provided with this paper.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript title and abstract promised an interesting and creative study, combining mechanistic RNA biology with developmental biology. However, while the study may have promise within the RNA field (although JUL had already been described previously), the authors clearly are not developmental biologists. The study is in this regard flawed in many ways. This applies to the introduction, experimental design, as well as data interpretation.\n\nA few examples of errors and misstatements in the introduction:\nLine 40-44: “This pattern suggests that vascular tissues, beyond resource transport, acted as structural and developmental axes, potentially serving as organizing centers for the establishment of leaf polarity and patterning. Developmental evidence supports this link: showing that the asymmetric distribution of specific molecular components establishes polarity, providing positional cues that guide pattern formation during organogenesis 5.”\nThe reference cited reviews the role of asymmetric cell divisions in generating distinct cell identities through cell division. There is no evidence for the vasculature as an organizing center for adaxial-abaxial leaf polarity.\nLine 45: In early leaf development, primary adaxial–abaxial polarity is first established6–9.\nIn fact, the cited work shows that adaxial-abaxial polarity is established before the formation of a vascular strand in the new primordium, contradicting the above premise.\nLine 50-51: Subsequently, surrounding tissues align with this vascular axis, specifying and differentiating to mesophyll and epidermis10,15.\nThe epidermis is specified during embryogenesis, and is maintained as a separate lineage throughout plant development.\nThe authors also confuse pri-miRNA and pre-miRNA. Pri-miRNA is the long primary transcript generated, in this case, by PolII. The shorter hairpin RNA, as shown in Fig. 1c, is the pre-miRNA.\nThe qRT-PCR analysis to monitor expression levels of different MIR165/166 genes and other polarity markers was performed on 4 week-old leaves. After 4 weeks, leaves are fully grown and patterned, any transcript differences observed are non-informative for developmental patterning. These experiments should be done on immature leaves, or vegetative apices. Given that expression is shown relative to WT, it is not clear whether any relevant transcript levels are detected.\nThe method section does not state how expression was normalized. In line 124, the authors state “We used pri-MIR166c and pri-MIR166d as internal negative controls that lack the putative minimal JUL1-binding sequence”. This would be the wrong choice. If expressed in 4 week-old leaves, these genes would be abaxial markers, and thus their expression is expected to change in the mutant, if not directly in an indirect manner via its effect on adaxial-abaxial polarity. A housekeeping gene that is unaffected in jul1jul2 mutants is needed to normalize expression across samples. The same applies for all other qRT analyses.\nLine 129-130: “To further assess the functional connection between JUL and miR165/166 in establishing the leaf adaxial–abaxial axis”. See the above point, the data presented does NOT address the establishment of adaxial-abaxial polarity. The authors make statements like this throughout the manuscript, but analysis of the establishment of leaf polarity requires studying the shoot apical meristem. Moreover, the phenotypes reported for jul1 and jul1 jul2 are far too mild to be caused by defects in leaf polarity establishment. The authors confuse “a minor effect on leaf polarity” with the far more exciting term “leaf polarity establishment”.\nThe phenotypes of the proRPS5A:STTM165/166 lines are surprisingly weak. Short Tandem Target Mimics, when constructed correctly, cause a dramatic decrease in the level of the targeted miRNA. As a result, STTM lines effectively targeting miR165/166 (when viable) show severe developmental defects, and changes in the expression of polarity related genes (e.g. Yan et al., 2012 doi: 10.1105/tpc.111.094144). The explanation provided by the authors on lines 146-148 does not make sense at all. This would indicate that miR165/166 has no function in leaf development. This raises questions about expression or effectiveness of the proRPS5A:STTM165/166 transgene in the lines selected.\nRegarding the vascular phenotypes shown in Extended data Fig. 3, the authors previously showed that JUL1 suppresses phloem development through its negative effects on SMXL4 and SMXL5. This complicates data interpretation. Are the observed defects due to effects on adaxial-abaxial polarity, lost regulation of SMXL genes, both, or even not yet discovered targets.\nLine 174: the term “emerging” is wrongly used\nLine 177: The authors conclude that JUL1-GFP accumulates throughout the leaf primordium. The quality of the data shown is not good enough to conclude this. The optical sections show many cells without GFP. Imaging actual leaf sections would generate a much greater resolution. Also, how many lines were analyzed? In other words, how reproducible is the data?\nThe tissue- or region-specific complementation of jul1jul2 is nice idea. However, how do the authors explain that expression of JUL1 on the adaxial side (BOP1 promoter) rescues the leaf curling defect of jul1 jul2? The latter is proposed to be caused by elevated miR165/166 levels in the mutant due to increased miRNA processing. However, the miR165/166 precursors are not expressed adaxially. This would point to an entirely different basis for the leaf curling phenotype.\nLine 199: see above. The data presented does not support a role for JUL1 in adaxial-abaxial pattern formation. The mutants show a mild leaf curling, potentially consistent with an effect on polarity maintenance.\nEvaluation of the RNA binding experiments are outside my expertise. However, two thoughts: 1) the on-bead binding assays are not reflective of what happens in the cell. The pri-miRNA transcript is able to form a hairpin structure, such that JUL1 has to compete with secondary structure formation. How realistic is that? Is single stranded pri-miRNA present, or can JUL compete with RNA folding? 2) The colocalization of JUL1 with MS2-tagged miR165a precursor is again not at all convincing. The resolution of this experiment is insufficient. MS2-GFP protein on its own is nuclear localized and, at the resolution provided, co-localizes with mRFP-JUL1 (left panel Fig 3e). The authors need to look at the sub-nuclear level. The statement that mRFP-JUL1(RA) does not co-localize with MS2-GFP is also not supported by the data. It is just that expression of mRFP-JUL1(RA) is comparatively low.\nAs is evident, I looked at the manuscript carefully up to the middle of page 9. However, I stopped reading at that point, as it was overwhelmingly clear that the current manuscript is not suitable for publication. The authors would be advised to consult a developmental biologist, and to greatly improve the quality of the data. Moreover, given that the authors identified over 65,000 regions capable of forming RG4s, they would need to think of additional experiments to convincingly make the point that increased miR165/166 levels are the cause, rather than an outcome of the jul1 jul2 phenotype. As it stands the study falls far short of its promise, there is no evidence for the model proposed.", "reviewer_2": "In this manuscript, Hwang and colleagues revealed new molecular and physiological functions of JUL1 in addition to regulating phloem development (Nature plants, 2018). They identify pri-MIR165/166 as new targets of JUL1 and uncover the regulatory role of JUL1 in miRNA processing, therefore provide new insights into the understanding of JUL1’s molecular functions. They also provided solid genetic and physiological proof that JUL1regulates leaf development dependent on MIR165/166 processing, therefore coordinates vascular development and leaf patterning, and update our understanding of coordinated development of plant tissues. Overall, this manuscript provides new insights into leaf development from RNA regulatory’s side with high quality data.\nMajor points:\n1) The authors have found JUL1 is differentially distributed in different cells, and this distribution seems to be very important for JUL1’s function and subsequently its regulatory role in leaf development, however the driving force of JUL1’s distribution is not clear. The authors may make some efforts to explore the factors that affect JUL1’s distribution, and the evolutionary selection of such factors across different species with different leaf shapes.\n2) Considering the RG4 formation, canonical RG4 sequence should include at least 4 G-blocks. However, in the pri-MIR165/166 sequence, UUGAGGGGAAUGUUGUCUGGAUCGAGGAU, only 3 G-blocks are found. Although there are 4G in the first block, it still cannot form two separate G-blocks. This sequence doesn’t satisfy the sequence requirement for RG4 formation. Given the complexity of RNA folding, there may be possibility of RG4 formation as indicated in Extended Data Fig 8 through circular dichroism (CD) spectra. However, to confirm RG4 formation, other methods should be included.\nMinor points:\n1) P should be always italicized and capitalized, such as in figure legend Figure 3 f.\n2) In reference, “Elife” should be in “eLife”."}, "author_response": "Dear editor,\nWe sincerely appreciate the reviewers’ time and thoughtful efforts in evaluating our manuscript. Their\ncomments have helped us substantially improve the clarity and rigor of the work, and we have addressed\nthe scientific concerns raised as carefully and thoroughly as possible in the revised version.\nHowever, we would like to respectfully raise a concern regarding the tone and balance of one of the\nreviewer’s comments. While we fully welcome critical and constructive feedback, certain remarks\nappear to reflect a degree of bias and a lack of objectivity in assessing the manuscript.\nFor instance, the reviewer states that “the authors clearly are not developmental biologists” and refers\nto the study as being “flawed in many ways.” We would like to clarify that, although some of the authors\nare trained primarily in RNA biology, others are developmental biologists with specific expertise in leaf\npolarity and vascular development. The overarching goal of this study was to integrate these\ncomplementary areas to explore the non-cell-autonomous role of JUL1 in modulating miR165/166\nactivity, as a coordinator between vascular development and leaf patterning, an interdisciplinary\nquestion that we believe is scientifically meaningful.\nWe acknowledge that a few developmental terms (e.g., “establishment” or “emerging”) were used\nimprecisely in the original version. These instances have been revised to align with conventional\ndevelopmental biology usage. However, we respectfully submit that these semantic issues do not\ncompromise the main findings or interpretations. Throughout the manuscript, we clearly present JUL1\nas a post-patterning regulator that contributes to the refinement or recalibration of polarity, not its initial\nestablishment in leaf development. This distinction is made explicit in both the revised Introduction and\nDiscussion sections.\nWe were especially concerned by the reviewer’s final comment: “I stopped reading at that point, as it\nwas overwhelmingly clear that the current manuscript is not suitable for publication.” While we\nrecognize that not all reviewers may agree with the conclusions or approach of a manuscript, we feel\nthat discontinuing the review midway and issuing a blanket rejection without a full evaluation of the\ndata limits the opportunity for fair and balanced assessment.\nWe appreciate the reviewer’s suggestion to consult a developmental biologist, and we emphasize that\ndevelopmental expertise, specifically in leaf polarity and vascular development, was integral to both\nthe design and interpretation of this study. We believe this perspective strengthens the interdisciplinary\nnature of our work, even if it was not fully recognized in the review. We believe interdisciplinary efforts,\nespecially at the interface of RNA biology and development, deserve rigorous but open-minded\nevaluation.\nWe hope that the revised manuscript, which incorporates extensive clarifications and new supporting\ndata, will be considered on its scientific merits. For your reference, please find our point-by-point\nresponses to the reviewers' comments below.\nBest regards,\nIldoo\nReviewers' comments:\nReviewer #1 (Remarks to the Author):\nThe manuscript title and abstract promised an interesting and creative study, combining\nmechanistic RNA biology with developmental biology. However, while the study may have\npromise within the RNA field (although JUL had already been described previously), the authors\nclearly are not developmental biologists. The study is in this regard flawed in many ways. This\napplies to the introduction, experimental design, as well as data interpretation.\nA few examples of errors and misstatements in the introduction:\nLine 40-44: “This pattern suggests that vascular tissues, beyond resource transport, acted as\nstructural and developmental axes, potentially serving as organizing centers for the\nestablishment of leaf polarity and patterning. Developmental evidence supports this link:\nshowing that the asymmetric distribution of specific molecular components establishes polarity,\nproviding positional cues that guide pattern formation during organogenesis 5.”\nThe reference cited reviews the role of asymmetric cell divisions in generating distinct cell\nidentities through cell division. There is no evidence for the vasculature as an organizing center\nfor adaxial-abaxial leaf polarity.\nAuthor response\nWe thank the reviewer for this insightful comment. We acknowledge that the cited reference (Ref.\n5) primarily discusses the role of asymmetric cell division in establishing distinct cell identities\nthrough specific molecular mechanisms at the cellular and tissue levels, and does not directly support\nthe notion that vascular tissues act as organizing centers for adaxial–abaxial polarity.\nOur intention was not to imply that vascular strands initiate polarity establishment. Rather, we aimed\nto propose that vascular development may contribute to the spatial refinement or coordination of\npolarity-related patterning during leaf development, based on currently identified molecular\nmechanisms mediating intercellular and intertissue communication.\nTo support this hypothesis, we refer not solely to Ref. 5, but to a growing body of genetic and\nphenotypic evidence demonstrating functional interplay between leaf polarity regulators and\nvascular patterning, including:\n Gain-of-function of the adaxial factor REVOLUTA (REV) disrupts miR165/166-mediated\ncleavage, leading to ectopic xylem arrangement surrounding the phloem (Emery et al., 2003,\nCurr. Biol.).\n Loss-of-function of KANADI1, an abaxial regulator, results in improper vascular bundle\nformation (Izhaki and Bowman, 2007, Plant Cell).\n Mutants of ASYMMETRIC LEAVES2 (AS2) and RH10, involved in adaxial identity and RNA\nregulation, exhibit abaxialized leaves with xylem dissipation (Matsumura et al., 2016, Biol.\nOpen).\n The hca2 mutant shows enhanced cambial activity and vascular proliferation along with curly\nleaf formation (Guo et al., 2009, Plant Cell).\n Overexpression of BREVIS RADIX (BRX), a regulator of phloem differentiation, leads to adaxial–\nabaxial patterning defects (Zhang et al., 2021, Hortic. Res.).\nThese findings collectively suggest that leaf polarity and vascular development are tightly\ncoordinated, likely via shared regulatory modules or non-cell-autonomous signaling pathways. We\nregret that this intended context may not have been sufficiently clear in the original text, and we\nhave revised the relevant passage to more accurately reflect this interpretation and avoid\nmisrepresentation.\nLine 45: In early leaf development, primary adaxial–abaxial polarity is first established6–9.\nIn fact, the cited work shows that adaxial-abaxial polarity is established before the formation of\na vascular strand in the new primordium, contradicting the above premise.\nAuthor response\nWe appreciate the reviewer’s insightful comment. We agree that our use of the term “establishment”\nmay have been imprecise and potentially misleading in the context of developmental biology, where\nit specifically refers to the initial specification of adaxial–abaxial polarity.\nIt is well established that leaf polarity is first initiated at the shoot apical meristem, prior to vascular\nstrand formation, and that vascular development is subsequently guided by this pre-established\npolarity. In fact, we describe this polarity-driven patterning process in the following sections of the\nmanuscript (Lines 45-52) and in the discussion (Lines 353-362).\nOur data do not suggest that vascular tissues serve as primary organizers of polarity establishment,\nbut rather to propose that vascular development may reinforce or spatially coordinate polarity cues\nduring later stages of leaf development. In our study, we interpret the role of JUL1 as that of a non-\ncell-autonomous coordinator, expressed in the vasculature, that contributes to the “refinement” or\n“recalibration” of leaf polarity as development progresses.\nTo avoid confusion and to better reflect our intended meaning, we have revised the relevant passage\nto replace “establishment” with a more appropriate term (Line 37-39).\nLine 50-51: Subsequently, surrounding tissues align with this vascular axis, specifying and\ndifferentiating to mesophyll and epidermis10,15.\nThe epidermis is specified during embryogenesis, and is maintained as a separate lineage\nthroughout plant development.\nAuthor response\nWe appreciate the reviewer’s correction and recognize the inaccuracy in our original statement. In\nour original wording, we unintentionally conflated the specification of the epidermal layer itself with\nthe specification and differentiation of epidermal cell types such as pavement cells, trichomes and\nguard cells, which acquire their identities during leaf development after initiation of vascular\ndevelopment.\nTo avoid this confusion, we have revised the sentence as follows: “Subsequently, surrounding tissues\nalign with this vascular axis, undergoing spatial organization and differentiation.” (Lines 48–50)\nThe authors also confuse pri-miRNA and pre-miRNA. Pri-miRNA is the long primary transcript\ngenerated, in this case, by PolII. The shorter hairpin RNA, as shown in Fig. 1c, is the pre-miRNA.\nAuthor response\nWe thank the reviewer for this comment. We would like to clarify that the structure shown in Fig.\n1c corresponds to the predicted secondary structure of the full-length pri-MIR165a, not the pre-\nmiRNA. To generate this structure, we used the published pri-MIR165a sequence (Zielezinski et al.,\n2015, BMC Plant Biology) and predicted its RNA folding using the RNAfold web server\n(http://rna.tbi.univie.ac.at//cgi-bin/RNAWebSuite/RNAfold.cgi). We hope this clarifies that pri-\nmiRNA and pre-miRNA were not conflated in our analysis.\nThe qRT-PCR analysis to monitor expression levels of different MIR165/166 genes and other\npolarity markers was performed on 4 week-old leaves. After 4 weeks, leaves are fully grown and\npatterned, any transcript differences observed are non-informative for developmental patterning.\nThese experiments should be done on immature leaves, or vegetative apices. Given that\nexpression is shown relative to WT, it is not clear whether any relevant transcript levels are\ndetected.\nAuthor response\nWe appreciate the reviewer’s important comment and apologize for the lack of clarity in our original\ndescription. We agree that fully expanded 4-week-old leaves are not ideal for assessing\ntranscriptional programs underlying developmental patterning.\nOur intention in analyzing these samples was not to infer causal relationships in early patterning, but\nrather to validate the phenotypic outcomes by examining whether the altered adaxial–abaxial ratios\nin mature leaves correlate with changes in the expression of established polarity marker genes (Fig.\n1 g and h and Fig. 2 f and g).\nTo specifically assess the role of JULGI in regulating primary and mature miR165/166 expression\nduring early developmental stages, we performed qRT-PCR using 7-day-old seedlings, which\ncontain the shoot apical meristem, leaf primordia, and developing leaves (Fig. 1 c).\nWe revised the Figure legends and Methods section accordingly to clarify the sampling stages and\nexperimental rationale (Line 522-523).\nThe method section does not state how expression was normalized. In line 124, the authors state\n“We used pri-MIR166c and pri-MIR166d as internal negative controls that lack the putative\nminimal JUL1-binding sequence”. This would be the wrong choice. If expressed in 4 week-old\nleaves, these genes would be abaxial markers, and thus their expression is expected to change in\nthe mutant, if not directly in an indirect manner via its effect on adaxial-abaxial polarity. A\nhousekeeping gene that is unaffected in jul1jul2 mutants is needed to normalize expression\nacross samples. The same applies for all other qRT analyses.\nAuthor response\nWe thank the reviewer for this important comment and fully acknowledge the concern regarding the\nuse of pri-MIR166c and pri-MIR166d as internal negative controls. As pointed out, these genes are\nexpressed in the abaxial domain and could be indirectly influenced in jul1 or jul1 jul2 mutants due\nto altered adaxial–abaxial polarity, especially in mature leaves.\nTo address this issue, we measured the expression of primary and mature miR165/166 transcripts in\n7-day-old seedlings, which contain only immature leaves and leaf primordia. This developmental\nstage minimizes potential secondary effects on abaxial gene expression. For all qRT-PCR analyses,\nwe normalized transcript levels to UBQ1, a housekeeping gene with stable expression that is\nunaffected in jul1 and jul1 jul2 mutants. Under these conditions, we observed no significant change\nin the expression of pri-MIR166c and pri-MIR166d, which lack the putative JUL1-binding sequence.\nIn contrast, pri-MIR165a and pri-MIR166a, which contain this motif, showed marked reductions in\ntranscript levels in both jul1 and jul1 jul2 mutants (Fig. 1d). These results support that pri-MIR166c\nand pri-MIR166d are not transcriptionally affected in these backgrounds under the tested conditions,\nthereby validating their use as non-target comparators. The differential expression patterns further\nsupport the specificity of JUL1-mediated transcriptional regulation.\nWe apologize for the confusion caused by the wording in line 124 and have revised the the Figure\nlegends and Methods section to clearly describe the normalization strategy used in all qRT-PCR\nexperiments (Line 522-523).\nLine 129-130: “To further assess the functional connection between JUL and miR165/166 in\nestablishing the leaf adaxial–abaxial axis”. See the above point, the data presented does NOT\naddress the establishment of adaxial-abaxial polarity. The authors make statements like this\nthroughout the manuscript, but analysis of the establishment of leaf polarity requires studying\nthe shoot apical meristem. Moreover, the phenotypes reported for jul1 and jul1 jul2 are far too\nmild to be caused by defects in leaf polarity establishment. The authors confuse “a minor effect\non leaf polarity” with the far more exciting term “leaf polarity establishment”.\nAuthor response\nWe appreciate the reviewer’s comment and acknowledge that our original wording was imprecise.\nAs noted above, we do not intend to suggest that JUL1 plays a role in the initial establishment of\nadaxial–abaxial polarity at the shoot apical meristem. Rather, our data support the view that JUL1\ncontributes to the modulation or refinement of polarity-related gene expression during later stages\nof leaf development, likely in coordination with vascular patterning (see Lines 353–362).\nWe recognize that the use of the term “establishment” may have overstated the functional\nimplications and caused confusion. To better reflect the scope of our findings and avoid\nmisinterpretation, we have revised the sentence to read:\n“To further assess the functional connection between JUL and miR165/166 in leaf adaxial–abaxial\npatterning, …” (Lines 129–130).\nThe phenotypes of the proRPS5A:STTM165/166 lines are surprisingly weak. Short Tandem Target\nMimics, when constructed correctly, cause a dramatic decrease in the level of the targeted miRNA.\nAs a result, STTM lines effectively targeting miR165/166 (when viable) show severe\ndevelopmental defects, and changes in the expression of polarity related genes (e.g. Yan et al.,\n2012 doi: 10.1105/tpc.111.094144). The explanation provided by the authors on lines 146-148\ndoes not make sense at all. This would indicate that miR165/166 has no function in leaf\ndevelopment. This raises questions about expression or effectiveness of the\nproRPS5A:STTM165/166 transgene in the lines selected.\nAuthor response\nWe thank the reviewer for this insightful comment. We agree that functional STTM constructs\ntargeting miR165/166 can cause dramatic phenotypes when expressed under strong, constitutive\npromoters, as shown in the study by Yan et al. (2012, Plant Cell). In that work, STTM165/166 was\ndriven by the 35S promoter, resulting in robust suppression of miR165/166 and severe\ndevelopmental defects.\nIn our study, we used the same STTM165/166 sequence design as described by Yan et al. (Rebuttal\nFigure 1). To validate the functionality of our construct, we expressed STTM165/166 under the 35S\npromoter and observed phenotypes consistent with strong miR165/166 depletion, thereby\nconfirming that the construct is effective when expressed at high levels.\nFor our experimental lines, we intentionally placed STTM165/166 under the control of the RPS5A\npromoter, which is active in the meristem and vascular initials but\nis less ubiquitous and potent than 35S. We hypothesize that this\nmore spatially and temporally restricted expression, along with [Figure Redacted]\nlower promoter strength, leads to partial not complete suppression\nof miR165/166, thereby producing milder but still detectable\nphenotypes. Rebuttal Figure 1. Phenotype of\nSTTM165/166 under the 35S promoter\nTo directly assess knockdown efficiency, we measured mature\nmiR165/166 levels in the shoot apices of 7-day-old seedlings. These data are now included in Figure\n1e–i and show a modest but reproducible reduction of miR165/166 (Fig. 1i), accompanied by\nmoderate de-repression of target genes. Correspondingly, phenotypes were present but not as severe\nas those in 35S lines.\nWe would like to emphasize that these results do not imply miR165/166 is dispensable for leaf\ndevelopment. Rather, they reflect the limited knockdown achieved with RPS5A-driven expression\nof STTM165/166. Importantly, our genetic interaction analyses show that the leaf phenotypes of\njul1 jul2 mutants are dependent on miR165/166 function, reinforcing the biological relevance of this\nregulatory relationship.\nRegarding the vascular phenotypes shown in Extended data Fig. 3, the authors previously showed\nthat JUL1 suppresses phloem development through its negative effects on SMXL4 and SMXL5.\nThis complicates data interpretation. Are the observed defects due to effects on adaxial-abaxial\npolarity, lost regulation of SMXL genes, both, or even not yet discovered targets.\nAuthor response\nWe appreciate the reviewer’s important point and fully agree that multiple regulatory layers, both\nknown and potentially unidentified targets of JUL1, may contribute to the leaf phenotypes observed\nin jul1 jul2 mutants.\nAmong the known targets, SMXL4 and SMXL5 are expressed specifically in phloem lineage cells,\nand their protein accumulation is similarly restricted to the phloem (Cho et al., 2018, Nature Plants;\nWallner et al., 2017, Current Biology). Given this spatial restriction, their direct involvement in\nestablishing or modifying adaxial–abaxial leaf polarity is unlikely. Consistent with this, smxl4 smxl5\ndouble mutants do not exhibit any defects in leaf polarity (Wallner et al., 2017, Current Biology).\nTo further evaluate this relationship, we examined plants carrying a targeted deletion of the JUL1-\nbinding site in the 5′ UTR of SMXL5 (Park et al., 2025, PNAS). These lines displayed an increase in\nphloem cell number, supporting the role of JUL1 in repressing SMXL5-mediated phloem\ndevelopment. However, they did not exhibit the leaf curling phenotypes observed in jul1 jul2\nmutants. Together, these findings suggest that the leaf patterning phenotypes are not mediated by\nSMXL4/5, and instead implicate a distinct regulatory mechanism.\nIn support of this, our biochemical data show that JUL1 directly binds to pri-miR165/166 and\nmodulates their processing, and that JUL1 protein is broadly distributed beyond the phloem. These\nobservations support a model in which JUL1 functions as a coordinator of both vascular\ndevelopment and leaf patterning, likely through independent pathways rather than solely via\nrepression of SMXL genes.\nLine 174: the term “emerging” is wrongly used\nAuthor response\nWe appreciate the reviewer’s comment and acknowledge that our original use of the term “emerging”\nwas imprecise. Our intent was not to suggest that the leaf was physically emerging from the shoot\napical meristem, but rather to refer to an early stage of leaf development. We agree that the term\n“emerging” may be misleading in the context of developmental biology. To clarify and avoid\noverstatement, we have revised the sentence as follows:\n“We detected GFP fluorescence indicative of MIR165a transcriptional activity in the abaxial domain\nof young leaf primordia in proMIR165a:GUS-GFP transgenic seedlings.” (Lines 174–175)\nLine 177: The authors conclude that JUL1-GFP accumulates throughout the leaf primordium. The\nquality of the data shown is not good enough to conclude this. The optical sections show many\ncells without GFP. Imaging actual leaf sections would generate a much greater resolution. Also,\nhow many lines were analyzed? In other words, how reproducible is the data?\nAuthor response\nWe appreciate the reviewer’s comment regarding the JUL1-GFP localization data. To assess\nreproducibility, we analyzed two independent proJUL1:JUL1-GFP transgenic lines and examined\napproximately 15 individual seedlings per line. In all cases, we observed a consistent pattern of\nJUL1-GFP distribution that was broader than that of the corresponding transcriptional reporter.\nThat said, we fully acknowledge that the resolution of the current confocal optical sections may not\nbe sufficient to support the phrasing “throughout the leaf primordium.” In particular, we recognize\nthat GFP signal may appear uneven due to optical interference from cellular structures such as\nchloroplasts, which can obscure fluorescence detection. In response, we have revised the manuscript\ntext to avoid overstatement and more accurately describe the observed expression pattern.\nImportantly, we note that JUL1-GFP signal is detectable even in abaxial epidermal cells of the leaf\nprimordium, where pri-MIR165a is expressed (Fig. 2a, b), supporting our conclusion that JUL1 may\ninfluence miR165/166 biogenesis in a non-cell-autonomous manner.\nThe tissue- or region-specific complementation of jul1jul2 is nice idea. However, how do the\nauthors explain that expression of JUL1 on the adaxial side (BOP1 promoter) rescues the leaf\ncurling defect of jul1 jul2? The latter is proposed to be caused by elevated miR165/166 levels in\nthe mutant due to increased miRNA processing. However, the miR165/166 precursors are not\nexpressed adaxially. This would point to an entirely different basis for the leaf curling phenotype.\nAuthor response\nWe thank the reviewer for raising this important point. We acknowledge that pri-MIR165/166\ntranscripts are primarily expressed in the abaxial domain of the leaf. However, in our study, we\nobserved that JUL1-GFP protein is distributed beyond its transcriptional origin, suggesting that\nJUL1 may function in a non-cell-autonomous manner.\nTo test this possibility, we performed tissue-specific complementation of the jul1 jul2 mutant using\nthe adaxial-specific BOP1 promoter. If JUL1 acted strictly in a cell-autonomous fashion, adaxial\nexpression would not be expected to affect miR165/166 processing in the abaxial domain or restore\nthe mutant phenotype. However, the observed rescue of the leaf curling phenotype by BOP1-driven\nJUL1 expression suggests that JUL1 can function non-cell-autonomously, potentially by influencing\ndownstream processes such as the movement or stability of pri-miRNAs, or through indirect\nregulatory effects across tissue layers.\nThese findings support the interpretation that JUL1, although expressed in specific tissues, can\ninfluence leaf development through non-cell-autonomous mechanisms. This is consistent with the\nbroader distribution of JUL1-GFP protein observed in our localization assays (Fig. 2a).\nLine 199: see above. The data presented does not support a role for JUL1 in adaxial-abaxial\npattern formation. The mutants show a mild leaf curling, potentially consistent with an effect on\npolarity maintenance.\nAuthor response\nWe respectfully note that our manuscript does not propose a role for JUL1 in the establishment of\nadaxial–abaxial polarity.\nAs described in the discussion part (line 358-364), JUL1 is expressed following the onset of phloem\ndevelopment and acts to modulate the polar distribution of the miR165/166–HD-ZIP III module.\nWe specifically use terms such as “recalibrate” and “feedback” to reflect a function in refinement or\nmaintenance, rather than in the initial establishment of polarity. We apologize if the phrasing\nelsewhere in the manuscript caused any confusion, and we have made minor revisions to the\nintroduction and results to clarify this distinction (Line 405‒409).\nEvaluation of the RNA binding experiments are outside my expertise. However, two thoughts: 1)\nthe on-bead binding assays are not reflective of what happens in the cell. The pri-miRNA\ntranscript is able to form a hairpin structure, such that JUL1 has to compete with secondary\nstructure formation. How realistic is that? Is single stranded pri-miRNA present, or can JUL\ncompete with RNA folding?\nAuthor response\nWe thank the reviewer for raising this important point. We fully agree that in vitro on-bead binding\nassays do not fully recapitulate the structural and dynamic context of pri-miRNA folding in vivo,\nwhere RNA secondary structures may limit access by RNA-binding proteins.\nTo address this limitation, we performed RNA immunoprecipitation (RIP) using jul1\nproJUL1:JUL1-HA complementation lines to evaluate the association between JUL1 and pri-\nMIR165a under native cellular conditions (Fig. 3f). While RIP cannot directly determine whether\nJUL1 competes with pri-miRNA secondary structures or preferentially binds to single-stranded\nregions, the enrichment of pri-MIR165a in JUL1-IP samples supports that JUL1 interacts with the\ntranscript in a physiological context.\nIn addition, using a protoplast transient expression system, we observed that JUL1 reduced the levels\nof RNA bound to the double-stranded RNA-binding domain of DCL1 (Fig. 4c), suggesting that\nJUL1 can affect DCL1-mediated pri-miRNA processing.\nThus, although we acknowledge the structural complexity of pri-miRNAs, our combined in vitro,\nprotoplast, and in planta RIP data collectively support JUL1 binding to pri-MIR165a in vivo and\ndemonstrate its potential impact on DCL1-dependent miRNA biogenesis.\n2) The colocalization of JUL1 with MS2-tagged miR165a precursor is again not at all convincing.\nThe resolution of this experiment is insufficient. MS2-GFP protein on its own is nuclear localized\nand, at the resolution provided, co-localizes with mRFP-JUL1 (left panel Fig 3e). The authors need\nto look at the sub-nuclear level. The statement that mRFP-JUL1(RA) does not co-localize with\nMS2-GFP is also not supported by the data. It is just that expression of mRFP-JUL1(RA) is\ncomparatively low.\nAuthor response\nWe thank the reviewer for the critical feedback. We acknowledge that the imaging resolution in the\ncurrent dataset is not sufficient to conclusively demonstrate subnuclear colocalization between\nmRFP-JUL1 and MS2-tagged pri-miR165a. We also agree that, given the broad nuclear localization\nof MS2-GFP, its apparent overlap with mRFP-JUL1 at the resolution presented may reflect co-\noccupancy of the nuclear compartment rather than true molecular colocalization.\nRegarding the JUL1(RA) mutant, we agree with the reviewer that the apparent lack of colocalization\nwith MS2-GFP may be confounded by lower expression levels of mRFP-JUL1(RA), rather than\nreflecting a biologically meaningful loss of interaction.\nTo address these concerns, using an Airyscan-equipped LSM800 microscope, we obtained higher-\nresolution images that revealed MS2-GFP colocalization with distinct cytosolic compartments\nassociated with mRFP-JUL1 in an RNA probe–dependent manner. Quantitative fluorescence\nintensity measurements within these condensates further confirmed probe-dependent enrichment of\nMS2-GFP signals, providing stronger evidence of colocalization (Fig. 3e).\nAs is evident, I looked at the manuscript carefully up to the middle of page 9. However, I stopped\nreading at that point, as it was overwhelmingly clear that the current manuscript is not suitable\nfor publication. The authors would be advised to consult a developmental biologist, and to greatly\nimprove the quality of the data. Moreover, given that the authors identified over 65,000 regions\ncapable of forming RG4s, they would need to think of additional experiments to convincingly\nmake the point that increased miR165/166 levels are the cause, rather than an outcome of the\njul1 jul2 phenotype. As it stands the study falls far short of its promise, there is no evidence for\nthe model proposed.\nReviewer #2 (Remarks to the Author):\nIn this manuscript, Hwang and colleagues revealed new molecular and physiological functions\nof JUL1 in addition to regulating phloem development (Nature plants, 2018). They identify pri-\nMIR165/166 as new targets of JUL1 and uncover the regulatory role of JUL1 in miRNA processing,\ntherefore provide new insights into the understanding of JUL1’s molecular functions. They also\nprovided solid genetic and physiological proof that JUL1regulates leaf development dependent\non MIR165/166 processing, therefore coordinates vascular development and leaf patterning, and\nupdate our understanding of coordinated development of plant tissues. Overall, this manuscript\nprovides new insights into leaf development from RNA regulatory’s side with high quality data.\nMajor points:\n1) The authors have found JUL1 is differentially distributed in different cells, and this distribution\nseems to be very important for JUL1’s function and subsequently its regulatory role in leaf\ndevelopment, however the driving force of JUL1’s distribution is not clear. The authors may make\nsome efforts to explore the factors that affect JUL1’s distribution, and the evolutionary selection\nof such factors across different species with different leaf shapes.\nAuthor response\nWe appreciate the reviewer’s insightful comment. We agree that the cell type–specific distribution\nof JUL1 is likely to be critical for its regulatory function in leaf development. Our observations\nindicate that JUL1 transcription is primarily confined to vascular tissues, while JUL1 protein forms\na gradient-like pattern that extends into surrounding cells. Given the size of JUL1-GFP (~42 kDa),\nwhich falls within the range permissive for plasmodesmal transport, we consider that passive\ndiffusion may contribute to establishing this distribution.\nAt the same time, we recognize that additional mechanisms—such as selective retention, degradation,\nor active transport—could also play important roles. While the present study was not designed to\ndissect the underlying drivers of JUL1 mobility, we agree this is an important direction for future\nresearch. In particular, identifying upstream factors that regulate JUL1 distribution will provide\nvaluable molecular insights into how its non-cell-autonomous function is controlled.\nWith respect to the evolutionary context, JULGI homologs are broadly conserved across vascular\nplants and are also found in certain non-vascular ancestral species (Park et al., 2025, PNAS). These\nfindings suggest that the functional establishment of JULGI is closely linked with the evolution of\nvascular tissues, since the vascular-derived, non-cell-autonomous feedback regulation we propose\npresupposes the presence of vasculature. Consistent with this idea, we observed defects in leaf\npatterning in other plant systems such as tomato and tobacco, supporting conservation of this\nmechanism across vascular plants to refine both vascular development and organ polarity. Ongoing\nstudies in our group aim to elucidate the ancestral roles of JULGI prior to the emergence of vascular\ntissues, which we expect will provide more direct insights into the evolutionary trajectory raised by\nthe reviewer.\n2) Considering the RG4 formation, canonical RG4 sequence should include at least 4 G-blocks.\nHowever, in the pri-MIR165/166 sequence, UUGAGGGGAAUGUUGUCUGGAUCGAGGAU, only 3\nG-blocks are found. Although there are 4G in the first block, it still cannot form two separate G-\nblocks. This sequence doesn’t satisfy the sequence requirement for RG4 formation. Given the\ncomplexity of RNA folding, there may be possibility of RG4 formation as indicated in Extended\nData Fig 8 through circular dichroism (CD) spectra. However, to confirm RG4 formation, other\nmethods should be included.\nAuthor response\nWe appreciate this insightful comment. We were aware of the limitation that the pri-MIR165/166\nsequence contains only three G-blocks and thus does not meet the canonical sequence requirement\nfor RG4 formation, which typically includes at least four G-blocks (5′-G≥3N1-7G≥3N1-7G≥3N1-7G≥3-3′). Indeed, the prediction scores obtained from in silico analyses were also lower compared\nto canonical RG4 sequences. For this reason, we did not interpret the observed features as definitive\nevidence of RG4 formation or directly link them to the molecular mechanism. Rather, we used this\nprediction solely as a reference criterion to guide the identification of potential JULGI targets. In\naddition, we performed CD spectral analysis to confirm that the selected sequence, although based\non RG4 prediction, behaved consistently during analysis, thereby validating the integrity of the\nexperimental procedure.\nFor further confirmation of RG4 formation in this sequence, we performed RT-STOP assays using\ntemplates containing either the wild-type G-rich elements or mutated elements of pri-MIR165a\n(Rebuttal figure 2). We observed a stop pattern immediately upstream of the last G-block under\npotassium or potassium/PDS (an RG4 stabilizer) treatment, indicating intra-molecular RG4\nformation. However, stops were predominantly detected near the beginning of the first and second\nG-blocks. Importantly, full-length transcripts remained the major products compared to the stalling\nproducts detected by RT-STOP. These results support the reviewer’s point that pri-MIR165/166\nlikely has a reduced capacity to form stable intra-molecular RG4 structures, while also raising the\npossibility that inter-molecular RG4 formation\nmight occur under certain conditions.\nIn line with the reviewer’s suggestion, we have now\nexplicitly stated in the revised manuscript that the G-\nrich elements in pri-MIR165a deviate from the\ncanonical RG4 sequence and therefore likely have a\nlower probability of forming stable intra-molecular\nRG4 structures. The revised text reads as follows:\n“We confirmed experimentally that pri-MIR165a\npotentially forms RG4 structures by in vitro circular\ndichroism (CD) analysis (Supplemental Figure 7).\nRebuttal figure 2. RT-STOP assay of G-rich elements in pri-\nHowever, considering the canonical RG4 forming MIR165a\nsequence, (5′-G≥3N1-7G≥3N1-7G≥3N1-7G≥3 -3′),\nthe G-rich elements in pri-MIR165a seem to have a lower probability of forming an RG4 structure.”\nMinor points:\n1) P should be always italicized and capitalized, such as in figure legend Figure 3 f.\nAuthor response\nWe have revised the manuscript so that P is consistently italicized and capitalized throughout,\nincluding in Figure 3f and all other figure legends.\n2) In reference, “Elife” should be in “eLife”.\nAuthor response\nWe have corrected the journal name “Elife” to the proper format “eLife” in the reference list."}, {"round": 2, "reviewer_comments": {"reviewer_2": "Thanks for the authors' effort answering all questions, no more questions are raised.", "reviewer_3": "Overview\nThis is an interesting study exploring a new contributor to the adaxial-abaxial polarity network: the RNA binding protein JULGI. I was invited to evaluate the author’s response to Reviewer 1 who had numerous critiques of the paper. These critiques were conceptual as well as experimental in nature. Conceptually, the authors did not clearly delineate ‘establishment’ from ‘maintenance’ phases of leaf polarity. Experimentally, the reviewer made several concrete suggestions to improve their experiments. These included better tissue choices, imaging, and in planta experiments. I believe these critiques have merit.\nFor instance, establishment vs maintenance is an extremely important distinction, as a lot of effort has gone into carefully defining the relative contributions of each polarity determinant to each phase in the process. Reading their revision, the authors have taken this critique to heart and changed most of the language to better reflect the ‘maintenance’ role of JULGI. However, a few lines still need to be modified:\nLine 62 states miR166 ‘establishes’ adaxial identity. However, Burian et al 2022 shows that AS2 and KAN are polarized across the ad-ab axis well before leaf emergence. The role of 166 is to limit HDZIPIIIs to the adaxial side; this is important because HDZIPIII factors are sufficient to convert abaxial identity into adaxial identity. However, HDZIPIII accumulation appears to be a response to – rather than the cause of – a polarized ad-ab axis. Line 363 has the same issue.\nLine 71 states vascular development ‘coordinates leaf patterning’. It is more accurate to say it ‘contributes to leaf shape’. Or that it ‘mediates/interprets underlying ad-ab polarity to impact leaf shape’. However they choose to write it, the text must make clear that patterning itself is not set up by JULGI or the vasculature.\nThe legend of Fig 5 still says ‘establishes’. I believe this is the last instance, but the text must be fully checked for removal/clarification of this issue.\nRegarding the experimental critiques, I did not see the original manuscript. However, the justifications laid out by the reviewer all seemed reasonable to me. Importantly, almost all these critiques have been addressed by the authors (see Weaknesses for two exceptions).\nStrengths\nMy own assessment of this version of the manuscript is that it is experimentally strong and conceptually interesting. Factors that aid in interpreting the underlying ad-ab axis are important and understudied. Given the importance of ncRNAs to ad-ab polarity, taking an RNA biology approach is also appropriate and interesting.\nThe experimental rigor in this version of the manuscript is outstanding. Indeed, multiple orthogonal assays are used to test each claim. I believe it is worth laying them out to clearly illustrate the number of independent assays performed:\n• To support a link between JUL1/2 and miR166 accumulation/activity: qRT for polarity determinants, STTM and qRT/phenotypic analyses\n• To support non-cell-autonomy: single cell data mining, transcriptional reporter, multiple translational reporters\n• To support binding of JUL1 to G-rich region of pri-MIR165a: REMSAs, bead-surface binding, circular dichroism, bacteriophage M2, in planta RIP\n• To support the notion that JUL1 prevents formation of dsRNA: bead surface binding, FRET\n• To support the notion that JUL1 prevents DCL1 from interacting with pri-MIR165A to process it into mir165: in planta RIP, pri-miRNA processing assay (see Weaknesses).\n• To support the notion that JUL1/2 act in the immediate term to suppress miR165a accumulation: protoplast co-transfection (see Weaknesses), short term induced JUL1 O/E and KO lines, MIR165amut phenotypic/qRT analyses in Wt and jul1/2 mutants\nI have concerns about a couple of these assays (see Weaknesses) but not enough to question their overall conclusions.\nWeaknesses\nThe pri-miRNA processing assay does not convincingly show a difference in mature 165 accumulation with JUL1 vs JUL1(RA), i.e. Fig 4d. Thus, conclusions that JUL1 works by inhibiting DCL1 processing are not supported. However, statements that JUL1 inhibits DCL1 interaction with pri-miR165A remain supported.\nThe RT-PCR for pri and mature 165A in Fig 5A does not show increases in either as claimed in text. JUL1 (and mir166 binding site)-dependent effects on GFP signal (via confocal) and protein levels (via Western) are supported, however. Thus, the only claims that can be made for Fig 5A are: 1) the presence of an intact miR165a binding site reduces accumulation of GFP protein, and 2) this reduction can be blocked via increased dosage of JUL1. The statement “These results suggest that JUL1 binding to pri-MIR165a suppresses the accumulation of mature miR165/166, leading to higher levels of transcripts for HD-ZIP III genes, the targets of miR165/166.” should thus be revised as these assays do not support this claim.\nMinor Comments and Suggestions\nLines 210, 214, 232 reference the wrong Extended Fig.\nThe authors should mention AGO10/PINHEAD/ZWILLE in the Intro or Discussion. AGO10 is a non-catalytic, adaxially-localized soak for miR166 (Zhu et al., 2011 Cell). As JULGI is an orthogonal post-transcriptional mechanism to limit 166 activity, a discussion of AGO10 in this context would be of interest.\nIn addition, AS2 should mentioned. There are many reasons to do this, including its involvement in the ad-ab pre-patterning (Burian et al) and the fact that it directly represses MIR166A (Husbands et al., 2015 Plant Cell)."}, "author_response": "Author response\nWe thank the reviewer for this insightful comment. We fully agree with the reviewer’s point, and have\nrevised the relevant sentences accordingly:\n(Line 61) In leaf development, abaxially expressed microRNA165/166 (miR165/166) maintains abaxial\nidentity by restricting the activity of class III homeodomain leucine zipper (HD-ZIP III) transcription\nfactors.\n(Line 364) In the abaxial domain, miR165/166 maintains abaxial identity by repressing HD-ZIP III, the\nkey determinants of adaxial fate.\n(Line 70) However, the spatiotemporal mechanisms by which vascular development mediates\nunderlying adaxial-abaxial polarity to shape leaf development, particularly through ncRNA regulation\nto integrate vascular and adjacent tissues during leaf formation, remain largely unknown.\nFigure 5. JUL1 refines leaf adaxial-abaxial patterning by suppressing miR165/166 levels in the abaxial\ndomain.\nRegarding the experimental critiques, I did not see the original manuscript. However, the\njustifications laid out by the reviewer all seemed reasonable to me. Importantly, almost all these\ncritiques have been addressed by the authors (see Weaknesses for two exceptions).\nStrengths\nMy own assessment of this version of the manuscript is that it is experimentally strong and\nconceptually interesting. Factors that aid in interpreting the underlying ad-ab axis are important\nand understudied. Given the importance of ncRNAs to ad-ab polarity, taking an RNA biology\napproach is also appropriate and interesting.\nThe experimental rigor in this version of the manuscript is outstanding. Indeed, multiple\northogonal assays are used to test each claim. I believe it is worth laying them out to clearly\nillustrate the number of independent assays performed:\n• To support a link between JUL1/2 and miR166 accumulation/activity: qRT for polarity\ndeterminants, STTM and qRT/phenotypic analyses\n• To support non-cell-autonomy: single cell data mining, transcriptional reporter, multiple\ntranslational reporters\n• To support binding of JUL1 to G-rich region of pri-MIR165a: REMSAs, bead-surface binding,\ncircular dichroism, bacteriophage M2, in planta RIP\n• To support the notion that JUL1 prevents formation of dsRNA: bead surface binding, FRET\n• To support the notion that JUL1 prevents DCL1 from interacting with pri-MIR165A to process\nit into mir165: in planta RIP, pri-miRNA processing assay (see Weaknesses).\n• To support the notion that JUL1/2 act in the immediate term to suppress miR165a\naccumulation: protoplast co-transfection (see Weaknesses), short term induced JUL1 O/E and KO\nlines, MIR165amut phenotypic/qRT analyses in Wt and jul1/2 mutants\nI have concerns about a couple of these assays (see Weaknesses) but not enough to question\ntheir overall conclusions.\nWeaknesses\nThe pri-miRNA processing assay does not convincingly show a difference in mature 165\naccumulation with JUL1 vs JUL1(RA), i.e. Fig 4d. Thus, conclusions that JUL1 works by inhibiting\nDCL1 processing are not supported. However, statements that JUL1 inhibits DCL1 interaction\nwith pri-miR165A remain supported.\nAuthor response\nWe appreciate the reviewer’s thoughtful comment. The assay in Fig. 4d was not intended to compare\nJUL1 and JUL1(RA), but rather to test whether differences in pri-miRNA binding affinity affect DCL1\nprocessing efficiency. Specifically, pri-MIR165a_WT and pri-MIR165a_mut#3 differ in their binding\naffinity to JUL1, while retaining identical mature miR165a sequence. Across all assay conditions, JUL1\naddition consistently reduced mature miR165a accumulation relative to reactions without JUL1, and\npri-MIR165a_mut#3, the weaker binder, produced higher mature miR165a levels, supporting a binding-\ndependent inhibitory effect on DCL1-mediated processing.\nWe agree, however, that our original wording could be misread as implying direct inhibition of DCL1’s\ncatalytic activity. We have revised the text to clarify that JUL1 interferes with pri-miRNA incorporation\ninto the DCL1 dsRNA-binding domain rather than inhibiting DCL1 catalysis:\n(Line 294) “…indicating that JUL1 reduces the efficiency of DCL1-mediated processing of pri-\nMIR165a into mature miR165. Collectively, these results suggest that JUL1 interferes with the\nincorporation of pri-MIR165a into the dsRNA-binding domain of DCL1, possibly through competitive\nbinding or steric hindrance, thereby suppressing miR165a biogenesis.”\nThe RT-PCR for pri and mature 165A in Fig 5A does not show increases in either as claimed in\ntext. JUL1 (and mir166 binding site)-dependent effects on GFP signal (via confocal) and protein\nlevels (via Western) are supported, however. Thus, the only claims that can be made for Fig 5A\nare: 1) the presence of an intact miR165a binding site reduces accumulation of GFP protein, and\n2) this reduction can be blocked via increased dosage of JUL1. The statement “These results\nsuggest that JUL1 binding to pri-MIR165a suppresses the accumulation of mature miR165/166,\nleading to higher levels of transcripts for HD-ZIP III genes, the targets of miR165/166.” should\nthus be revised as these assays do not support this claim.\nAuthor response\nWe thank the reviewer for this valuable observation. We agree that the visual differences in the RT-\nPCR bands in Fig. 5A appear subtle, although changes in band intensity are evident in a JUL1 dosage-\ndependent manner. To clarify this, we performed densitometric quantification of the pri- and mature\nmiRNA bands (Revision Figure 1, below). This analysis revealed an approximately 20% increase in\npri-miR165a and ~30% decrease in mature miR165/166 upon JUL1 expression, whereas no such\nchanges were detected with JUL1(RA). These quantitative results indicate that JUL1 modestly, but\nspecifically, affects pri-miR165a processing efficiency in this assay. However, as the reviewer noted,\nthis conclusion is not essential to support the main finding without the in planta data, and we have\ntherefore removed the conclusive statement from the text.\nRevision Figure 1. Quantification of RT-PCR bands in Fig. 5A\nImportantly, these data serve as supportive evidence consistent with the in planta results shown in Fig.\n1d and Fig. 5b,c, where reciprocal and statistically robust changes in pri- and mature miR165/166 were\nobserved in jul1/jul1 jul2 mutants, inducible JUL1 overexpression, and JUL1 knockdown lines.\nMinor Comments and Suggestions\nLines 210, 214, 232 reference the wrong Extended Fig.\nAuthor response\nWe thank the reviewer for carefully noting this error. The references in lines 210, 214, and 232 were\nindeed incorrect and should refer to Extended Data Fig. 8 rather than Extended Data Fig. 7. These have\nbeen corrected in the revised manuscript.\nThe authors should mention AGO10/PINHEAD/ZWILLE in the Intro or Discussion. AGO10 is a\nnon-catalytic, adaxially-localized soak for miR166 (Zhu et al., 2011 Cell). As JULGI is an\northogonal post-transcriptional mechanism to limit 166 activity, a discussion of AGO10 in this\ncontext would be of interest.\nIn addition, AS2 should mentioned. There are many reasons to do this, including its involvement\nin the ad-ab pre-patterning (Burian et al) and the fact that it directly represses MIR166A\n(Husbands et al., 2015 Plant Cell).\nAuthor response\nWe thank the reviewer for this insightful suggestion. We fully agree that AGO10/PINHEAD/ZWILLE\nand ASYMMETRIC LEAVES2 (AS2) are key components of the adaxial–abaxial regulatory\nframework and highly relevant for contextualizing the role of JUL1. In the revised manuscript, we have\nadded a discussion highlighting how AGO10, AS1/AS2, and HD-ZIP II maintain adaxial identity by\nlimiting miR165/166 expression or activity in the adaxial domain, and how JUL1 operates through a\nspatially distinct mechanism by modulating miR165/166 biogenesis in the abaxial domain.\nThis addition clarifies that JUL1 represents an orthogonal post-transcriptional layer that refines adaxial–\nabaxial balance rather than reinforcing adaxial suppression. We added this part as follows in Discussion,\n(Line 369) Mechanistically, this process parallels the action of ARGONAUTE10 (AGO10), as both\nlimit miR165/166 activity through post-transcriptional regulation58,59. Whereas AGO10,\nASYMMETRIC LEAVES1/260, and HD-ZIP II28 act in the adaxial domain to restrict miR165/166\naccumulation, JUL1 acts in the abaxial domain to modulate miR165/166 biogenesis. Thus, JUL1 fine-\ntunes the abaxial source strength of miR165/166, adding a distinct layer of spatial regulation to the\npolarity network."}]

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