S₁ basic leucine zipper transcription factors shape plant architecture by controlling C/N partitioning to apical and lateral organs

Significance

Most plants exhibit a pronounced apical growth dominance. However, when the resource demand of the primary organ decreases or the nutritional status of the plant increases, apical growth dominance is broken, and lateral organ growth unleashed. In our work, we have unraveled a developmental function of the sugar-controlled S₁-basic-leucine-zipper (S₁-bZIPs) transcription factors in mediating apical growth dominance, by controlling tissue-specific expression of genes involved in sugar (C)-partitioning or control of organic nitrogen-(N) homeostasis. By these means, S₁-bZIPs could act as rheostat to tune lateral organ outgrowth, by restricting their C/N-supply. Known to be repressed at high-sugar levels, S₁-bZIPs have the capacity to integrate information on resource availability into growth plasticity, which is of great relevance for breeding strategies.

Abstract

Plants tightly control growth of their lateral organs, which led to the concept of apical dominance. However, outgrowth of the dormant lateral primordia is sensitive to the plant’s nutritional status, resulting in an immense plasticity in plant architecture. While the impact of hormonal regulation on apical dominance is well characterized, the prime importance of sugar signaling to unleash lateral organ formation has just recently emerged. Here, we aimed to identify transcriptional regulators, which control the trade-off between growth of apical versus lateral organs. Making use of locally inducible gain-of-function as well as single and higher-order loss-of-function approaches of the sugar-responsive S₁-basic-leucine-zipper (S₁-bZIP) transcription factors, we disclosed their largely redundant function in establishing apical growth dominance. Consistently, comprehensive phenotypical and analytical studies of S₁-bZIP mutants show a clear shift of sugar and organic nitrogen (N) allocation from apical to lateral organs, coinciding with strong lateral organ outgrowth. Tissue-specific transcriptomics reveal specific clade III SWEET sugar transporters, crucial for long-distance sugar transport to apical sinks and the glutaminase GLUTAMINE AMIDO-TRANSFERASE 1_2.1, involved in N homeostasis, as direct S₁-bZIP targets, linking the architectural and metabolic mutant phenotypes to downstream gene regulation. Based on these results, we propose that S₁-bZIPs control carbohydrate (C) partitioning from source leaves to apical organs and tune systemic N supply to restrict lateral organ formation by C/N depletion. Knowledge of the underlying mechanisms controlling plant C/N partitioning is of pivotal importance for breeding strategies to generate plants with desired architectural and nutritional characteristics.

Plants have an enormous capacity to modulate their architecture. However, many agronomically important plant species as well as the model plant Arabidopsis (Arabidopsis thaliana) exhibit a growth dominance of the apical meristems, while the pre-established lateral shoot (axillary branch) and root (lateral root, LR) primordia remain initially dormant (1, 2). This control of nutrient-intensive growth of lateral organs in support of apical meristems, referred to as apical dominance, is beneficial for the plant’s Darwinian fitness (3). It prevents plants from being overgrown by neighboring plants competing for light, water, and minerals and ensures species conservation in fluctuating environments by concentrating limited resources on the primary inflorescence to allow seed maturation. As such, apical dominance provides a gateway for successful dispersal into competitive and resource-scarce habitats without the need for genetic change and is therefore a key determinant of plant evolutionary success.

Nevertheless, upon silique ripening and the concomitant decrease in carbohydrate (C) demand of the growing shoot tip, repression of axillary shoot outgrowth is released, allowing an upscaling of seed production (4). Consistently, decapitation of the primary shoot apex, which results in rapid C reallocation to axillary branches (5) or cultivation under high-light conditions that increase total C availability (6) strongly reduce apical dominance. Similar observations have been made for LR growth, which is induced by either root apical meristem excision (7), root sugar feeding, or high-light cultivation (8, 9), suggesting a coordinated, C-responsive expansion of shoot and root tissues (2, 9). Indeed, recent studies demonstrate that C redistribution to lateral organs represents the initial trigger for primordia outgrowth and is dominant over the repressive effects of the well-established branching-control hormones auxin and strigolactone (5, 10). Despite its pivotal importance, the underlying sensing and signaling mechanisms that enable plants to integrate nutritional inputs and direct resources to apical or lateral meristems to modulate overall architecture remain largely unresolved.

However, major advances in uncovering transporter activities that facilitate C fluxes from the site of production (source) to consuming tissues (sink), assist in revealing the coordination of C partitioning. For instance, sugar fluxes in Arabidopsis originate largely from photosynthetically active rosette leaves, which act as strong C source tissue. While a significant fraction of leaf C is stored as starch, sucrose can passively diffuse from the leaf mesophyll cells either across plasmodesmata to neighboring tissues or preferentially along the gradient to the apoplastic space via sucrose efflux transporter proteins (SUGARS WILL EVENTUALLY BE EXPORTED TRANSPORTERS, SWEETs) (11). For long-distance transport to distal sinks, incoming apoplastic sugars are actively loaded into the phloem companion cells by sucrose-H⁺ symporters of the SUCROSE TRANSPORTER family (SUC/SUTs) and are either unloaded symplastically or by SWEETs at the growing, C-demanding apical shoot, or root meristems (12). This creates a constant flow of sugars from the source tissue to the distal apical meristems, while the proximal lateral sinks remain C-depleted, thereby maintaining apical growth dominance.

In addition to C supply, the availability of organic N is essential for meristem growth (4). Most N uptake takes place in roots that absorb nitrate or ammonium via specialized membrane-localized transporters from the surrounding soil (13). While ammonium serves directly as a substrate for root GLUTAMINE SYNTHETASE (GS) isozymes to form glutamine (Gln) from glutamate, nitrate is primarily transported through the xylem to the leaves, where it also feeds the Gln pool after initial enzymatic reduction to ammonium and final conversion by leaf GS (13). Gln, the most abundant amino acid (AA) in plants, is used as a major transport form of organic N and acts as a central nitrogen donor for the biosynthesis of proteinogenic AAs or nucleotides for DNA replication. To control homeostasis of this key AA, Gln can be deaminated by glutaminases such as the recently identified GLUTAMINE AMIDO-TRANSFERASE 1_2.1 (GAT1_2.1) to initiate its degradation via the tricarboxylic acid cycle (14). Importantly, loss-of-function mutant analysis has implicated GAT1_2.1 in shoot branching control (15).

In plants, two highly conserved and counteracting metabolic kinases, namely TARGET OF RAPAMYCIN (TOR) and SUCROSE-NON-FERMENTING1 RELATED KINASE1 (SnRK1), are known to be central components of a C/N status integrating system that controls plant growth and metabolism by reprogramming global gene expression and enzymatic activities (16–18). Whereas TOR is activated by a high C/N ratio and promotes anabolism and plant growth responses, SnRK1 signaling is activated under energy-deficient conditions and induces catabolic processes and growth repression (17, 18). Although functionally well-characterized in managing severe energy deprivation, recent work highlights the impact of SnRK1 and its catalytic subunit SnRK1α1 in controlling sink tissue development, such as seedling establishment (19) and lateral root growth (20), respectively. Although SnRK1 addresses multiple targets in this respect, the downstream group C basic leucine zipper (bZIP) transcription factor (TF) bZIP63, which preferentially heterodimerizes with group S₁-bZIPs (S₁-basic-leucine-zipper) upon phosphorylation (21, 22), appears to implement key aspects of the SnRK1α1-driven transcriptional adaptation (19, 20, 23). Consistently, SnRK1α1 strongly boosts the transactivation potential of group S₁-bZIPs (24), leading to the concept of an interconnected SnRK1-C/S₁ bZIP heterodimerization network (25).

Along this line, recent work proposed the importance of individual members of the highly homologous S₁-bZIP TFs: bZIP1, bZIP2, bZIP11, bZIP44, and bZIP53 in mediating growth repression and metabolic adaptation to stress conditions (23, 26, 27). As they possess an additional layer of post-transcriptional regulation through conserved upstream open reading frames in their 5′ leader sequences that confer sucrose-induced repression of translation, it is postulated that they have the intrinsic capacity to integrate sucrose availability into growth responses (28). Although S₁-bZIPs exhibit highly overlapping expression domains in source (leaf petioles of mature leaves) and sink (young leaves, root tip, flowers, and rosette core with branches) (29) tissues, their functional implications have not been addressed, particularly due to limited availability of mutants and potential functional redundancy.

In this study, we focused our attention on unraveling the role of group S₁-bZIP TFs in controlling energy-intensive lateral organ formation. Therefore, we used CRISPR/Cas9 to generate single and higher-order mutant combinations of S₁-bZIPs and analyzed their growth throughout the vegetative and reproductive phases. In contrast to wild-type (WT) plants, which showed a marked dominance of apical shoot and root growth, we observed a clear promotion of lateral organ growth at the expense of apical organs in higher-order mutants of S₁-bZIPs, suggesting their importance for establishing apical dominance. Strikingly, removal of excessive C-demanding branches or increasing root C supply by sugar feeding rescued the impaired apical growth responses of the mutants. Therefore, we hypothesized that S₁-bZIPs are required for C-partitioning to apical meristems. Consistent with this hypothesis, mutants showed a strong reduction in sugar availability in the leaf apoplast and distal sinks, but an increase in proximal sinks (axillary rosette branches) and leaf cells, indicative of an impaired sugar export from the latter. Further metabolic analyses revealed an additional function of S₁-bZIPs in controlling root and shoot N (especially Gln) homeostasis. Applying comprehensive transcriptomics, molecular biology approaches, and mutant analysis, we identified specific clade III SWEETs and the glutaminase GAT1_2.1 as direct S₁ targets involved in controlling tissue-specific C and N supply. Therefore, we propose that the energy-controlled S₁-bZIP TFs act as key regulators of shoot and root apical dominance by determining the C/N status of apical and lateral meristems.

Results

Higher-Order Mutants of Group S₁-bZIP TFs Exhibit a Growth Repression of Apical and Growth Promotion of Lateral Organs.

In recent years, gain-of-function approaches implicated the low-energy responsive group S₁-bZIP TFs in mediating systemic growth repression (26, 30). Nevertheless, they were found to be expressed in specific C-sink and -source tissues under non-starved conditions, suggesting their involvement in sink-source communication and control of plant development (29). However, a lack of mutants and potential functional redundancy among the highly homologous S₁ members severely hampered their functional characterization. To assess the impact of group S₁ bZIP members in this respect, we used CRISPR/Cas9 (31) to generate individual single bZIP knockouts and higher-order mutant combinations, ranging from double to a full group S₁ quintuple mutant (SI Appendix, Table S1). While single and double mutants, e.g., of the phylogenetically closely related bZIP1 and bZIP53 showed no obvious alterations in vegetative growth (SI Appendix, Fig. S1A), a triple mutant of the close homologs bzip2, bzip11, and bzip44 (bzip2/11/44) or a full group S₁ quintuple mutant (bzip1/2/11/44/53) showed, after a WT-like expansion of the first true leaves, a slight (bzip2/11/44) to marked (bzip1/2/11/44/53) leaf growth retardation, respectively (Fig. 1 A and B). This mutant phenotype coincided with an impaired primary root growth response (Fig. 1 C and D) and surprisingly an increased lateral root emergence compared to the WT (Fig. 1E).

Fig. 1.

Mutants of S₁ bZIP TFs exhibit a growth repression of apical and growth promotion of lateral organs. (A) Shoot growth of S₁ bZIP mutants is impaired at late vegetative stage. Representative pictures of soil-grown 2-(Upper row) and 4-(Lower row) week-old WT, bZIP triple (bzip2/-11/-44), and quintuple (full group S₁, bzip1/-2/-11/-44/-53) mutants. (Scale bar: 1 cm.) (B) Mean leaf area (±SEM, shaded area, n = 24 per genotype) of WT (black line), triple (red line), and quintuple (blue line) bZIP mutants from 14 to 28 days after germination (DAG). (C) Primary root growth of S₁ bZIP mutants is impaired. Representative picture of 7-(Left) and 10-d-old (Right) WT, bZIP triple, and quintuple mutant seedlings. (Scale bar: 1 cm.) (D) Mean PR length (±SEM, n = 6, each replicate consisting of 10 plants per genotype) of WT (black line), bZIP triple (red line), and quintuple (blue line) seedlings was measured from 5 to 12 DAG. (E) Emerged lateral root density (eLRD) over time of the WT (black line), bZIP triple (red line), and quintuple (blue line) mutants. Given are mean values (±SEM, n = 6, each replicate consisting of 10 plants per genotype). (F) Bolting of S₁ bZIP mutants is delayed. Representative picture of the 6-wk-old WT, bZIP triple, and quintuple mutants. (Scale bar: 1 cm.) (G) Time of bolting of the WT (black box), bZIP triple (red box), and quintuple (blue box) mutants referred to the day after germination. Given are mean values (n = 16 to 24 per genotype). (H) Mean seed weight (n = 15 per genotype) of the WT (black box) and bZIP triple (red box) mutants. (I) Mean silique length (n = 7 to 8 per genotype) of siliques 6 to 15 (counted from the inflorescence base) from the 8-wk-old WT (black box) and bZIP triple (red box) mutants. (J) S₁ bZIP mutants exhibit a high branching phenotype. Representative picture of the 12-wk-old WT and bZIP quintuple mutants. (Scale bar: 1 cm.) Magnification shows a high number of extremely short and frequently aborted siliques. (Scale bar: 1 cm.) (K) Mean number (±SEM, n = 22 per genotype) of axillary rosette branches of the WT (black line) and bZIP triple (red line) or (L) WT (black line) and bZIP quintuple mutants (blue line) (±SEM, n = 13 to 16 per genotype). Outgrown axillary rosette branches (≥1 cm) were counted starting 5 days after bolting (DAB). (M) Local overexpression of bZIP11 in rosette cores represses axillary branching but does not affect primary shoot growth (SI Appendix, Fig. S1M). Representative picture of mock treated (solvent without dexamethasone, −DEX) and DEX induced (+DEX) Pro35S::bZIP11-GR plants at 7 days after start of induction (DAI). Treatment started prior to first axillary rosette branching when primary inflorescences reached ~10 cm. DEX or the solvent control was applied every second day to the rosette cores. Red arrows mark outgrown axillary rosette branches. (Scale bar: 1 cm.) (N) Mean number (±SEM, n = 24 per treatment) of outgrown axillary branches of Pro35S::bZIP11-GR plants after recurring mock (black line) or DEX (green line) treatment. (O) Schematic picture of differences in apical and lateral organ growth of the WT and S₁ bZIP mutants. Plants were either grown on soil (Fig. 1 A, B, and F–N) or on ½-strength MS medium (Fig. 1 C–E) under long day conditions at 100 µmol photons m⁻² s⁻¹. Statistically significant differences were determined by one-way ANOVA (Fig. 1 B, D, and E), Kruskal–Wallis ANOVA (Fig. 1G), Student’s t-test (Fig. 1 H and I), or Mann-Whitney U test (Fig. 1 K, L, and N) comparing the WT and respective bZIP mutants (***P < 0.001). For details, see Dataset S2.

Similar alterations in apical and lateral organ growth were observed during the reproductive phase. While bolting time of the primary inflorescence of the single and double mutants resembled that of the WT (SI Appendix, Fig. S1B), the triple, and quintuple bZIP mutants exhibited a moderate or substantial delay, respectively (Fig. 1 F and G). In addition, seed and silique growth at the primary inflorescence was significantly impaired in triple (Fig. 1 H and I) and severely affected in quintuple mutants, which showed hardly any viable siliques (Fig. 1J). Remarkably, silique growth of the triple mutant could be largely rescued by removing all resource-consuming axillary branches that were about to grow out (SI Appendix, Fig. S1C), indicating that silique growth was not generally defective in the mutant, but rather short in resource supply.

In contrast to the impaired apical shoot growth responses, growth of lateral shoot organs was enhanced. The bzip11 mutant, for instance, showed a mild and transient increase of axillary branching, while the other single and double mutants showed no obvious growth phenotype (SI Appendix, Fig. S1 D–H). In contrast, the triple (Fig. 1K) and individual quintuple mutants (Fig. 1 J and L and SI Appendix, Fig. S1I) exhibited an early and strong enhancement of shoot branching.

Based on the bzip11 branching phenotype and a strong bZIP11 induction in leaf petioles and axillary buds upon establishment of the primary inflorescence (SI Appendix, Fig. S1 J and K), we used this TF as a prototypical example for complementary gain-of-function approaches. To avoid artefacts caused by constitutive and systemic overexpression, we employed a dexamethasone (DEX)-inducible expression system, which allowed a rapid and tissue-specific translocation of bZIP11 into the nucleus (30). Whereas repeated DEX applications did not alter bud outgrowth of the WT (SI Appendix, Fig. S1L), it severely suppressed axillary branching of Pro35S:: bZIP11-GR plants (Fig. 1 M and N), while growth of the primary inflorescence remained unaffected (SI Appendix, Fig. S1M).

In conclusion, the comprehensive phenotypic characterization of the S₁-bZIP loss- and gain-of-function lines clearly demonstrates a key function of S₁-bZIPs in regulating apical and lateral shoot and root growth. Furthermore, a striking correlation between the order of S₁ mutations and the reduction in apical dominance suggests a largely redundant function of these TFs (Fig. 1O).

S₁-bZIP TFs Mutants Show a Strongly Reduced Leaf Sugar Export to the Apoplast for Long-Distance Transport to Distal Sinks.

The promotion of lateral shoot and root growth at the expense of the respective apical organs and the rescue of impaired silique growth by removal of lateral sinks (branches) pointed toward a shifted nutrient allocation between apical and lateral organs in the S₁-bZIP mutants. This was further supported by an increased C/N ratio in the rosettes and a slightly decreased C/N ratio in flowers of the triple mutants compared to the WT (SI Appendix, Fig. S2 A–C). To analyze C allocation in more depth, we examined the WT and bZIP mutants for their sugar content in primary roots and in apical and lateral shoot organs.

Concerning roots, we assessed C supply by determining the concentration of sucrose, the major sugar transport form, in primary roots of the WT, bZIP triple, and quintuple mutants. Strikingly, we found a 30 to 40% reduction in root sucrose levels in the mutants (Fig. 2A), supporting the notion that S₁-bZIPs could be involved in root sugar provision. To test whether a reduced sucrose supply caused the primary root growth repression and to rule out an irreversible impairment of primary root growth in the mutants, we performed sugar-feeding experiments. Remarkably, cultivation of the triple and quintuple bZIP mutants on half-strength MS media supplemented with 30 or 60 mM sucrose, largely restored primary root growth to WT levels (Fig. 2 B and C and SI Appendix, Fig. S2 D–F). Notably, external sucrose supply also restored LR formation in a sucrose concentration–dependent manner (SI Appendix, Fig. S2 G–O).

Fig. 2.

S₁ bZIP TFs are crucial for leaf sugar export to the apoplast for long-distance transport to distal sinks. (A) Primary roots of S₁ bZIP mutants are sucrose depleted. Given is the mean root sucrose content (n = 8 individual pools of roots per genotype) of 12-d-old WT (black box), bZIP triple (red box), and quintuple (blue box) mutants. (B and C) Sucrose feeding can largely revert impaired root growth of S₁ bZIP triple (red box) and quintuple (blue box) mutants back to the WT (black box) level. Plants were grown for 10 d on ½ strength MS medium supplemented with 30 or 60 mM of sucrose (Suc) and/or mannitol (Man) as osmotic control. (B) Representative picture of 7-d-old WT and bZIP mutant plants grown under control conditions or sucrose supplementation. (Scale bar: 1 cm.) (C) Given is the mean primary root length (bar box plot, n = 6, each replicate consisting of 10 plants per genotype) of the WT (black box), bZIP triple (red box), and quintuple (blue box) mutants in the presence or absence of sucrose. (D and E) Distal sinks of S₁ bZIP mutants are sugar depleted, while sugar supply of proximal sinks is not affected. (D) Glucose and (E) sucrose content (n = 7) was determined by UPLC–MS/MS in the 8-wk-old WT (black box) and bZIP triple mutants (red box). Stated tissues were harvested when primary inflorescence reached ~10 cm. (F) Starch levels were determined at ZT0 from flowers of the primary inflorescence of the WT (gray box) and triple mutants (red box) (n = 7) prior branching. (G) Leaf sugar export to the apoplast is strongly impaired in S₁ bZIP mutants. Sugar (sucrose, fructose, and glucose) content in AWF of the WT (black box) and bZIP triple mutants (red box) was determined by UPLC–MS/MS. Whole rosettes of 6-wk-old plants were vacuum-infiltrated with water and AWF was collected by centrifugation. Given are mean values (log₂, n = 9 to 10 per genotype) derived from AWF of two plants per replicate. (H and I) Leaves of S₁ bZIP mutants accumulate high levels of starch. (H) Mean leaf starch content (±SEM, n = 3 per genotype) throughout the day/night cycle (light/dark shaded area) of the WT (black line) and bZIP triple mutant (red line). (I) Mean starch content (n = 7, each replicate consisting of two plant rosettes) in rosettes leaves of the 5-wk-old bolted WT (black box), bZIP triple (red box), and quintuple (blue box) mutants at ZT0. Starch was determined enzymatically. (J) Relative CO₂ fixation rate of leaves of the 5-wk-old WT (gray line) and triple mutants (red line) under light or darkness (light/dark shaded area). Provided are mean values (±SEM, n = 4/5 per genotype). (K) Mean leaf sucrose levels (±SEM, n = 4 per genotype) throughout the day/night cycle (light/dark shaded area) of the WT (black line) and bZIP triple mutant (red line). (L and M) Leaf sugar content in DEX inducible Pro35S::bZIP11-GR plants. Starch (L) as well as sucrose, glucose, and fructose (M) levels were determined after a short-term (6 h) DEX (green boxes) or solvent-control (gray boxes) treatment (n = 6 per treatment). (N) Schematic picture of sugar levels in distinct sink or source tissues of S₁ bZIP mutants relative to the WT (red arrows). Sugar transport routes from source to sink tissues (black arrows) as well as localization of sugar export (red boxes) or import (green boxes) facilitators and cwINV are given. Plants were either grown on soil (Fig. 2 D–M) or ½-strength MS medium (Fig. 2 A–C) under long day conditions at 100 µmol photons m⁻² s⁻¹. Statistically significant differences were determined by one-way ANOVA (Fig. 2A), Mann-Whitney U test (Fig. 2 C, E, K, and M), or Student’s t-test (Fig. 2 D, F–J, and L) comparing the WT and respective bZIP mutants, n.s. not significant, *P < 0.05, **P < 0.01, ***P < 0.001. For details, see Dataset S2.

To define the sugar content in apical and lateral shoot organs (sinks) and shoot source tissues, we determined the levels of the two most abundant sugars—glucose (Fig. 2D) and sucrose (Fig. 2E) in mature leaves (source) as well as young leaves, rosette cores enriched in axillary rosette buds, and flowers (sinks). This was done at an early reproductive stage, prior branching to examine whether changes in sugar content might precede the observed growth alterations. As the quintuple mutant however showed extremely early and strong axillary branching right after inflorescence bolting, which affects global sugar levels, we excluded this mutant from the analysis and focused on the WT and triple mutant. While we did not observe any difference in glucose or sucrose levels in leaves of either genotype (Fig. 2 D and E), we detected a strong decrease in distal sink tissues (flowers) of the triple mutant (Fig. 2 D and E). Consistently, starch reserves in the flowers of the triple mutant were strongly reduced compared to the WT (Fig. 2F). In contrast, triple mutants showed a significant increase in glucose levels in proximal sinks (rosette core with axillary rosette buds) compared to WT plants (Fig. 2D). Therefore, we hypothesized that leaf sugar export is impaired in the triple mutant, which would increase the symplastic sugar flux to proximal sinks but inhibit long-distance transport through the apoplast and vasculature to distal sinks. To further pursue this, we isolated leaf apoplastic wash fluid (AWF) from whole rosette leaves and measured the concentration of sucrose and its degradation products glucose and fructose that are produced by apoplastic, cell wall–associated invertases (cwINV). Consistent with an impaired sugar export from mesophyll cells, apoplastic sugars were severely reduced by about 30% (for glucose) to 50% (for sucrose or fructose) in the mutant (Fig. 2G).

An impaired sugar export from leaf cells should ultimately favor starch biosynthesis to counteract the toxicity of high intracellular hexose concentrations. Accordingly, we found significantly elevated starch levels in the triple mutant (Fig. 2H), especially during the night and the beginning of day. A slightly stronger effect was also observed in the quintuple mutant at dawn (Fig. 2I). As high leaf starch levels were proposed to negatively feedback on photosynthesis (32), it is worth noting that the triple mutant showed a reduced CO₂ assimilation rate (Fig. 2J) and an impaired leaf sucrose accumulation over the day (Fig. 2K) compared to WT plants. In contrast to this, DEX-induced overexpression of bZIP11 in rosette leaves resulted in a significant reduction in leaf starch (Fig. 2L) and to an increase in sucrose, glucose, and fructose levels (Fig. 2M), indicative of an enhanced sugar export to the leaf apoplast. Collectively, we conclude from these results that group S₁-bZIP TFs operate in leaves to support sugar export to the apoplast and, consequently, long-distance transport to distal sinks, such as the apical shoot and root meristems (Fig. 2N).

Shoot and Root Gln Supply Is Controlled by S₁-bZIP TFs.

Besides C availability, supply of organic N is essential for meristem activity. Under low-energy conditions, individual group S₁-bZIPs were found to induce degradation of proline and the branched chain AAs leucine, isoleucine, and valine to feed carbon skeletons into alternative respiratory pathways for ATP production (23). To obtain a comprehensive picture of the organic N status during development of the higher-order bZIP mutants under non-starved conditions, we determined the levels of proteinogenic AAs in shoot and root tissues of the WT, bZIP triple, and quintuple mutants in the middle of the day at an early and rather late vegetative stage. In contrast to what was found in response to energy depletion, we did not observe a considerable change in proline and branched chain AA levels throughout tissue development of the genotypes (Fig. 3 A–C and SI Appendix, Fig. S3). However, a striking increase in the levels of the most abundant AAs Gln and asparagine (Asn), which serve as major organic N transport forms during the day or night, respectively (33), was detected in seedling roots of the triple (Fig. 3A) and, to an even greater extent, in the quintuple mutant (SI Appendix, Fig. S3) compared to the WT. Although the effect on transport AAs was not found in the shoot tissue of the triple mutant at the seedling stage (Fig. 3B), a clear and specific increase in Gln levels was observed just before bolting (Fig. 3C). In contrast, we observed at the respective stage a significant reduction of Gln and asparagine levels in rosette leaves of the Pro35S::bZIP11-GR line, shortly after DEX application (Fig. 3D). Therefore, we propose that group S₁-bZIP TFs redundantly act in roots to systemically control homeostasis of the central organic N transport form Gln during plant development.

Fig. 3.

S₁ bZIP mutants show a marked increase in Gln levels in roots and shoots prior bolting. (A) Given is the mean AA content (bar box plots, n = 8) determined by UPLC–MS/MS from roots of 2-wk-old WT and bZIP triple mutant seedlings. Root material of several plants was harvested at ZT8 and pooled. (B) Mean AA content (bar box plots, n = 8) in shoots of the 2-wk-old WT (black box) and bZIP triple mutant (red box) seedlings. Shoot material of several plants was harvested at ZT8 and pooled. (C) Mean AA content (bar box plots, n = 5, each replicate consisting of two plants) from rosettes of 4-wk-old WT or bZIP triple mutant plants harvested at ZT8. (D) Mean AA content (bar box plots, n = 6 per treatment) from rosettes of 4-wk-old DEX inducible Pro35S::bZIP11-GR plants after 6 h of DEX (green box) or solvent (gray box) treatment. Plants were grown on (A and B) ½ strength MS medium or (C and D) soil under long-day conditions at 100 µmol photons m⁻² s⁻¹. Statistically significant differences were determined by the Mann-Whitney U test comparing individual AA content of the WT and respective bZIP mutants (Dataset S2).

S₁-bZIP TFs Control Tissue-Specific Expression of Defined Clade III SWEET Sugar Transporters and of the Glutaminase GAT1_2.1.

To gain insight into the mechanisms by which S₁-bZIPs control local C/N supply, we performed RNA-sequencing (RNAseq) to identify differentially expressed genes (DEGs) in source and sink tissues of the triple and quintuple S₁-bZIP mutants. To visualize the complex dataset (Dataset S1), we generated Venn diagrams showing the significantly (P < 0.05) up- (log₂FC ≥ 1) or down-regulated (log₂FC ≤ −1) genes of the triple (Fig. 4 A and B) or quintuple (Fig. 4 C and D) mutant relative to WT plants. Consistent with previous work revealing individual S₁-bZIPs as transcriptional activators recruiting the histone acetylation machinery (34), we generally found more down- than upregulated genes in the triple and quintuple mutants (Fig. 4 A–D). In addition, we observed a high degree of tissue specificity, suggesting sink/source-specific regulation of S₁ factors. Focusing on genes that are involved in managing C distribution, we found among the DEGs several sugar transporters of the SWEET family, known to facilitate leaf sugar export (35), (Fig. 4 E and F). Of these, predominantly clade III SWEETs (SWEET9-15) (11) were affected, with SWEET10, -13, and -14 strongly down-regulated in sinks and SWEET15 down-regulated in both source and sink tissues (Fig. 4 E–G). To address whether S₁-bZIPs have the potential to directly bind and control their expression, we analyzed a publicly available DNA affinity purification sequencing (DAPseq) database (36) to examine direct, in vitro promoter binding. In addition, we performed an assay for transposase-accessible chromatin by sequencing (ATACseq) (37) from protoplasts of transgenic plants that were transfected with a DEX-inducible bZIP11 construct and were treated with the translation inhibitor cycloheximide to uncover direct, in vivo bZIP11-dependent chromatin opening of SWEET promoter regions. While we observed only a weak binding of S₁-bZIPs to the promoter of SWEET10 and -13 (SI Appendix, Fig. S4 A and B) associated with marginal chromatin opening (SI Appendix, Fig. S4 E and F), we found a strong binding (SI Appendix, Fig. S4 C and D) and site-specific bZIP11-dependent chromatin opening for the SWEET14 and especially SWEET15 promoter (SI Appendix, Fig. S4 G and H). Applying protoplast transactivation assays (PTAs), we validated the transactivation properties of S₁-bZIPs on the highly responsive SWEET15 promoter (Fig. 4H) and pinpointed two cognate bZIP cis-elements (Gbox-related elements, GREs) as critical for promoter transactivation (Fig. 4I). Moreover, we found SWEET15 to be transcriptionally induced in rosette leaves of the DEX induced Pro35S::bZIP11-GR line (Fig. 4J).

Fig. 4.

S₁ bZIP TFs control expression of SWEET15 and GAT1_2.1 in distinct plant tissues. (A–D) DEGs between the WT and S₁ bZIP mutants in distinct C source and sink tissues. Global transcript abundance in specific tissues (axillary rosette buds, blue; open flower, red; young leaves, green; and old leaves, yellow) of the WT, bZIP triple, and quintuple mutant plants at the pre-branching state was determined by RNA sequencing (n = 3 per genotype and tissue). Venn diagrams present significantly (P < 0.05) Up (log₂FC ≥ 1) (A and C) or Down (log₂FC ≤ −1) (B and D) regulated genes comparing the WT, bZIP triple, or quintuple mutant plants, respectively. The complete list of DEGS can be found in Dataset S1. (E and F) Overview of tissue specific expression of SWEET family members (grouped in clades I to IV) in (E) bZIP triple or (F) bZIP quintuple mutants relative to the WT. Given are log2-fold expression changes (induction given in shades of red, repression in shades of green). (G) Validation of SWEET10, -13, -14, and -15 expression by q-RT-PCR in flowers of the WT (black bar), bZIP triple (red bar), or quintuple (blue bar) mutants prior branching. Presented are mean expression values (±SEM, n = 3) relative to UBIQUITIN5 expression. (H and I) PTAs using a SWEET15 promoter–driven GUS reporter (ProSWEET15:GUS) and Pro35S-driven S₁ bZIP effector constructs. Pro35S-driven NAN expression was used for normalization. (H) SWEET15 promoter activity in the absence (−) or presence of bZIP effector constructs (bZIP1, -2, -11, -44, -53) given as mean GUS/NAN values (±SEM, n = 3) relative to promoter background levels. (I) Promoter activity of the original and GRE mutated SWEET15 promoter in the absence (−) or presence of bZIP11 effector construct given as mean GUS/NAN values (±SEM, n = 2) relative to promoter background levels. (J) Expression analysis of SWEET15 and GAT1_2.1 in rosettes of 4-wk-old Pro35S::bZIP11-GR plants after short-term (6 h) DEX (green bar) or solvent (black bar) treatment by q-RT-PCR. Presented are mean expression values (±SEM, n = 6/5) relative to UBIQUITIN5 expression and normalized to the solvent control. (K) Expression analysis of GAT1_2.1 in shoots and roots of the 2-wk-old WT (black bar), bZIP triple (red bar), or quintuple (blue bar) mutants by q-RT-PCR. Presented are mean expression values (±SEM, n = 3) relative to UBIQUITIN5 expression. (L and M) PTAs using a GAT1_2.1 promoter–driven GUS reporter (ProGAT1_2.1:GUS) and Pro35S-driven S₁ bZIP effector constructs. Pro35S-driven NAN expression was used for normalization. (L) GAT1_2.1 promoter activity in the absence (−) or presence of bZIP effector constructs (bZIP1, -2, -11, -44, -53) given as mean GUS/NAN values (±SEM, n = 3) relative to promoter background levels. (M) Promoter activity of the original and GRE mutated GAT1_2.1 promoter in the absence (−) or presence of bZIP11 effector construct given as mean GUS/NAN values (±SEM, n = 2) relative to promoter background levels. Plants were either grown on soil (A–J, L, and M) or ½-strength MS medium (K) under long day conditions at 100 µmol photons m⁻² s⁻¹. Statistically significant differences were determined by Student’s t-test comparing the WT and respective bZIP mutants, n.s. not significant, *P < 0.05, **P < 0.01, ***P < 0.001. For details, see Dataset S2.

To identify targets that are causative for the increased root Gln levels in S₁-bZIP mutants, we screened a previously published transcriptomics dataset of bZIP11 overexpression lines (30). Among the DEGs, the glutaminase GAT1_2.1 emerged as the most promising candidate as it 1) was strongly induced by bZIP11 in leaves (Fig. 4J) (30), 2) was predominantly expressed in roots (15), and 3) showed a broad binding of S_1-bZIPs in a GRE-rich promoter region (SI Appendix, Fig. S4I), which also exhibited a significant bZIP11-dependent increase in chromatin accessibility (SI Appendix, Fig. S4J). Consistent with these results, we found a strong downregulation of root GAT1_2.1 expression in the bZIP triple and, to an even greater extent, in the quintuple mutant (Fig. 4K). We also observed a strong and redundant activation of the GAT1_2.1 promoter by S₁-bZIPs (Fig. 4L), which was at least partially accomplished by two adjacent GREs located within the proposed bZIP11 binding site (Fig. 4M).

Mutant Lines Affecting S₁-bZIP Targets in C/N Allocation Exhibit Shoot Branching Phenotypes.

TFs generally control a plethora of target genes, which result in complex phenotypes. To assess whether the addressed clade III SWEETs and the GAT1_2.1 glutaminase contribute significantly to the S₁-bZIP-driven repression of lateral organ growth, we monitored axillary branching of loss-of-function mutants of the respective target genes. While we observed no increase in shoot branching between WT plants and an unrelated (clade II) sweet8 mutant (SI Appendix, Fig. S5A), mutants in clade III sweets such as sweet15 (Fig. 5 A and B), sweet10 and sweet13 (SI Appendix, Fig. S5 B and C) showed moderate branching phenotypes, while sweet14 mutations had only a very mild effect (SI Appendix, Fig. S5C). In line with previous reports (14, 15), we also observed a substantial increase in shoot branching of the gat1_2.1 mutant (Fig. 5 C and D). In conclusion, specific clade III SWEETs and GAT1_2.1 are well-suited targets to participate in S₁-bZIP mediated growth control of lateral organs.

Fig. 5.

Mutants of SWEET15 and GAT1_2.1 reveal moderate branching phenotypes. (A–D) Axillary rosette branching of SWEET15 and GAT1_2.1 mutants. (A) Representative picture of 6-wk-old WT and two independent sweet15 plants. Red arrows mark outgrown axillary rosette branches. (Scale bar: 1 cm.) (B) Mean axillary branching (±SEM, n = 24 per genotype) of WT (black line) or sweet15 (green line) plants. (C) Representative picture of 5-wk-old WT and gat1_2.1 plants. Red arrows mark outgrown (≥1 cm) axillary rosette branches. (Scale bar: 1 cm.) (D) Mean axillary branching (±SEM, n = 24 per genotype) of WT (black line) or gat1_2.1 (yellow line) plants. Plants were grown on soil under long day conditions at 100 µmol photons m⁻² s⁻¹. Outgrown axillary rosette branches (≥1 cm) were counted starting 5 DAB. Statistically significant differences were determined by Mann-Whitney U test comparing the WT and respective mutants (Dataset S2).

Discussion

Plants display an enormous developmental plasticity in shaping shoot and root architecture. Although most plants show a pronounced apical growth dominance, lateral organ formation is inducible and highly sensitive to the plant’s sugar status (6, 9). However, how metabolism and development are coordinated is far from being understood (38). Recent work demonstrated that rapid sugar reallocation to dormant axillary rosette branches upon decapitation acts as an initial trigger for bud outgrowth (5) and highlights the impact of sugar signaling on shoot branching (39–42). This suggests that plants possess a sophisticated mechanism for sensing nutrient availability and allocating limited resources between apical and lateral meristems to control their growth.

Initially characterized as central players in the plant’s energy management system (25), we unraveled in this study a developmental function of the subgroup S₁-bZIP TFs (bZIP1, -2, -11, -44, and -53) in tuning apical growth dominance. Known to be expressed under energy-depleted conditions (43) and in strong sink tissues [such as flowers (29), seeds (44), and roots (26)], we also observed bZIP11 expression in petioles of source leaves upon establishment of the primary inflorescence sink (SI Appendix, Fig. S1 J and K). This suggests that S₁-bZIP expression is controlled by either tissue-specific or developmental signals, if not both. This hypothesis is supported by the RNAseq datasets derived from source and sink tissues of the S₁-bZIP mutants (Fig. 4 A–D), which show a high degree of tissue specificity in bZIP-mediated expression. Unexpectedly, the typical starvation-responsive targets of S₁-bZIPs (e.g., ETFQO, MCCA, etc.) (23) are not differentially expressed in the analyzed tissues, suggesting a clear difference in bZIP action in starvation-related or developmental processes.

Consistent with the marked decrease in sugar supply to the leaf apoplast and apical sink tissues (Fig. 2 A and D–G), and sugar increase in leaf cells and proximal sinks such as the axillary rosette branches (Fig. 2 D, H, and I), we observed a shift from apical growth promotion to lateral organ formation in the S₁-bZIP mutants (Fig. 1 A–L). These correlative results pointed towards a coordinated, S₁-bZIP-dependent change in sugar supply of the apical sinks. In line with this and emphasizing that S₁-bZIP mutants are not generally defective in apical shoot and root growth, sugar supplementation of the C-depleted roots or reduction in C-demand of the shoot by excision of all axillary branches rescued the mutant’s apical growth phenotypes such as primary root growth or silique expansion on the primary shoot inflorescence (Fig. 2 B and C and SI Appendix, Fig. S1C). Examination of PTAs (Fig. 4 H and I), RNAseq (Fig. 4 E and F), ATACseq (37), and DAPseq (36) datasets (SI Appendix, Fig. S4 A–H) identified specific clade III SWEET sugar transporters as potential targets in this regard. Consistently, the single sweet10, sweet13, sweet14, and sweet15 as well as the double sweet13/-14 mutants showed a moderate reduction in apical dominance (Fig. 5 A–D and SI Appendix, Fig. S5 A–C), and the latter also had shortened siliques and increased lateral root formation (45), partially resembling S₁-bZIP mutant phenotypes (Fig. 1 I–L). Higher-order mutants of clade III SWEETs could further assist in unraveling their joint contribution to S₁-bZIP-driven apical dominance. However, due to prevalent SWEET heterodimerization (46), these mutants would likely exhibit highly pleiotropic effects. While the discussed SWEETs are co-ordinately downregulated in sink tissues of the bZIP mutants, SWEET15 expression was also affected in leaves, likely leading to impairment of both sugar export from the source and import at apical sink tissues (Fig. 4 E and F) and consequently reduced apical growth. In line with this, we observed in leaves of the triple mutant an accumulation of starch (Fig. 2H) and a reduction of sucrose levels (Fig. 2K), while DEX-induced bZIP11 overexpression in leaves led to SWEET15 induction (Fig. 4J), a drop in leaf starch (Fig. 2L) and an increase in leaf sucrose, glucose, and fructose (Fig. 2M), which pointed towards an improved apoplastic transport.

Importantly, both SWEET expression and sugar translocation to apical meristems are equally impaired in bZIP triple and quintuple mutants, indicating that bZIP2, -11 and -44 are particularly crucial in this respect (Figs. 2 A and I and 4G). In contrast to the strongly down-regulated SWEETs, SWEET8, which is central for pollen development (47) as well as SWEET11 and -12, which have been proposed to act in tandem to facilitate phloem loading at source and unloading at sink tissues (48) are strongly upregulated in floral tissues of the bzip triple and/or quintuple mutants (Fig. 4 E and F). Since S₁-bZIPs are well described as transcriptional activators, the observed effects are likely to be indirect and may represent a compensatory response to reduced sugar availability in mutant flowers.

In addition to their function in C partitioning, we also uncovered the impact of S₁-bZIPs on Gln homeostasis. Representing an important reservoir for organic N, Gln is essential for organ growth and development. In this respect, we observed a clear correlation between the order of S₁-bZIP mutations, the increase in root Gln levels (Fig. 3A and SI Appendix, Fig. S3) and the expression of lateral organ growth (Fig. 1 E, K, and L and SI Appendix, Fig. S2 G, J, and M). It is worth noting that Gln levels in the mutants were only elevated in roots and not in shoot tissues at the early vegetative stage (Fig. 3 A and B), while a strong increase was observed in the shoot at the onset of the reproductive phase (Fig. 3C). This suggests that either root Gln is efficiently transported to the shoot upon bolting and/or that again bZIP action is subject to tissue- and development-specific regulation. Making use of PTAs (Fig. 4 L and M), transcript analysis by qRT-PCR (Fig. 4 J and K), ATACseq (37), and DAPseq (36) data (SI Appendix, Fig. S4 I and J), we identified the recently described glutaminase GAT1_2.1 (14) as a direct target of S₁-bZIPs. Consistently, largely overlapping expression domains of GAT1_2.1 and S₁-bZIPs were revealed in sink tissues, such as flowers, siliques, axillary rosette branches, and roots (15, 29). In line with this, all S₁ factors appear to contribute to the full expression of GAT1_2.1, which is most severely downregulated in the quintuple S₁ mutant (Fig. 4K). Complementary to the loss-of-function approach, inducible bZIP11 overexpressors showed a strong and rapid activation of GAT1_2.1 transcription, which was accompanied by a marked decrease in plant Gln levels (Figs. 3D and 4 J) (30). Furthermore, DEX-application to the rosette core of this line, revealed a strong bZIP-dependent repression of axillary branching (Fig. 1 M and N). Conclusively, phenotypic analysis of a gat1_2.1 mutant validated its strong impact on shoot branching (15) (Fig. 5 C and D).

Notably, the strength of the branching phenotype correlated with the order of S₁-bZIP mutations (Fig. 1 K and L) and Gln levels (Fig. 3 A–C and SI Appendix, Fig. S3) as well as reduction in GAT1_2.1 expression (Fig. 4K). In contrast, the effects on sugar allocation (Fig. 2 A and I) and SWEET expression (Fig. 4G) were comparable between the triple and quintuple bzip mutants, suggesting differences in gene regulation and that modulation of N levels might act as a prominent factor for S₁-bZIP dependent branching control.

In summary, we propose that S₁-bZIPs have a dual function in establishing apical growth dominance, by controlling C and N allocation to apical and lateral organs (Fig. 6). Based on our results, we suggest that this is accomplished by S₁-bZIP TFs controlling SWEET expression in source (leaf petioles) and distal sink tissues (shoot and root apical meristems) to generate a constant sugar flow to the growing apical organs, while keeping the lateral primordia in a sugar depleted state. The central impact of S₁-bZIPs on root glutaminase abundance and in consequence global organic N supply further assists in tuning organ development.

Fig. 6.

Working model of S₁ bZIP action in C and N partitioning to control shoot and root architecture. The model plant Arabidopsis as well as many agronomically important plant species display a pronounced apical growth dominance, while lateral organs remain quiescent until the sink demand of the primary meristem declines. To enable such a differential growth control of apical shoot/root meristems and the lateral shoot (axillary rosette branch)/root (lateral root) primordia, plants require a mechanism to direct fluxes of C (sugars, black arrows) and N resources (AAs, yellow arrow) from places of production to the specific resource-consuming organs that are destined to grow (sink tissues). This is accomplished by coordinated sugar and AA partitioning. While sucrose efflux transporter family proteins (SWEETs) facilitate passive diffusion of photosynthesis-derived sugars from rosette leaves to the leaf apoplast, sugar transporters of the SUC/SUTs family actively load apoplastic sugars against the gradient into the phloem. Predominantly at strong apical sinks, like the flowering shoot apex or the growing primary root meristem, SWEETs again assist in unloading of phloem sugars, thereby creating a constant sugar gradient toward the apical meristems. In reverse orientation, Gln, which serves as a transport form of root-fixed N is loaded into the root xylem to be transported to the actively growing shoot and root sinks. As S₁ bZIPs directly and redundantly control expression of central clade III SWEETs at source (leaf) and sink (flower) tissues and of root GLUTAMINE AMIDOTRANSFERASE 1_2.1 (GAT1_2.1) that is known to initiate Gln degradation, they potentially control C partitioning to the primary meristems and balance systemic N homeostasis. Consistently, higher-order mutants of S₁ bZIPs are disturbed in long-distance sugar transport and display enhanced leaf starch accumulation and diffusion of hexoses, likely via symplastic routes, to proximal sinks, such as axillary rosette branches and lateral roots. An additional strongly impaired GAT1_2.1 expression in the mutants, results in a marked increase in shoot and root glutamine (N) supply providing all resources required for lateral meristem outgrowth. In conclusion, S₁ bZIPs have the capability to manage C/N partitioning to promote apical but restrict lateral meristem activity.

As S₁-bZIPs are highly conserved in agronomically relevant plants, manipulation of this system would help to create plants with increased or reduced branch number, which would be valuable for fruit, flower, and timber industries. In addition, local misexpression could be used to direct sugar fluxes to specific tissues. Consistently, recent reports have shown that fruit-specific misexpression of S₁-bZIP orthologs in strawberry or tomato results in a strong increase in fruit sugar content (49–51). Thus, our study broadens the view on the developmental impact of the stress-related S₁-bZIP TFs to direct C/N allocation to control energy-intensive plant architecture, thereby opening a gateway for biotechnological progress.

Material and Methods

Transgenic Lines, Plant Cultivation, and Phenotypic Characterization.

In all assays, WT A. thaliana accession Columbia (Col-0) was used as control, and all transgenic lines in this study are in a Col-0 background. The Pro35S::bZIP11-GR line was characterized by Hanson and co-workers (30). For induction, 10 µM DEX (Merck, Germany) dissolved in DMSO was used. The sweet13 (SALK_087791, psi00187), sweet14 (SALK_010224, psi00188), and sweet13 x sweet14 (psi00189) mutants were characterized previously (45) and were obtained from RIKEN (https://www.riken.jp). The sweet15 mutant line was characterized and kindly provided by the Coupland lab. The gat1_2.1 (SALK_031983C, N665686) mutant was characterized previously (14) and was obtained from NASC (https://arabidopsis.info/). ProbZIP11::bZIP11-GFP and ProbZIP11::bZIP11-GUS plant lines were constructed by PCR-amplifying 2.3 kb of the bZIP11 promoter and the joining bZIP11 coding region without stop from genomic DNA using Gateway®-site flanking primers, given in SI Appendix, Table S2. The sweet8, sweet10, and all single and higher-order mutants of S₁-bZIP TFs were generated in this work using an egg-cell-specific CRISPR/Cas9 genome editing system (31). Higher-order mutants were generated by a multiplexed CRISPR approach using arrays of target-specific guide-RNA expression cassettes (31). The quintuple (bzip1/2/11/44/53) mutant was created by transforming the Cas9 free bzip2/11/44 triple mutant with a multiplexed CRISPR construct specific for bZIP1 and bZIP53. Highly target-specific guide sequences (showing no predicted off-targets), which were positioned close to the translational start codon, ATG, were obtained from ChopChop (52) and are given in SI Appendix, Table S2. Transgenic lines were selected by hygromycin treatment. Homozygous mutants, exhibiting frame-shifts and consequently premature stop codons in SWEET and bZIP genes were identified by sequencing PCR products of the putatively mutated regions. Genotyping primers used for this step are listed in SI Appendix, Table S2. An overview of mutations in the respective mutants is given in SI Appendix, Table S1. A detailed description of generation of transgenic lines and determination of plant growth parameters can be found in SI Appendix.

Sugar Measurements.

Unless stated otherwise, sugar (glucose, fructose, sucrose) measurements were performed as published previously (20), with minor modifications. In brief, around 20 mg (fresh weight) of individual tissues (roots, leaves, rosette cores with branches or flowers) were harvested at ZT0 and frozen in liquid N₂. Sugars from ground tissue were extracted with 300 µL 80% (v/v) ethanol, containing 2 µg [1,1-²H₂]-trehalose and 8 µg [6,6-²H₂]-glucose as internal standards. Samples were incubated at 80 °C for 20 min and centrifuged for 10 min at 18,000 × g. After transferring the supernatant to a new reaction tube, the pellet was reextracted twice, first with 300 µL 50% (v/v) ethanol and, subsequently with 300 µL 80% (v/v) ethanol both at 80 °C for 20 min. The extracts were pooled, and the solvent completely evaporated using a vacuum concentrator at 45 °C. For MS (UPLC–MS/MS) analysis, the obtained pellet was redissolved in 25 µL 50% (v/v) methanol. Apoplastic sugars were extracted based on a previously published method (53) with minor modifications. A detailed protocol can be found in SI Appendix.

AA Measurements.

Around 50 mg (fresh weight) of shoot or root material of individual genotypes were frozen in liquid nitrogen and ground to powder. To each sample, 300 µL of extraction solvent [50% (v/v) MeOH, 1.07 mg/mL methyl methanethiosulfonate, 10 µM norvaline, 50 mM HCl] was added. Samples were shaken for 10 min at 21 Hz in a cooled mixer mill. After centrifugation at 4 °C and 18,000 × g for 10 min, 300 µL of the supernatant was transferred to a fresh tube. The pellet was then reextracted with 300 µL extraction solvent as before and the supernatants combined. After centrifugation (4 °C and 18,000 × g for 10 min), 20 µL of the final supernatant was then derivatized using the AccQ-Tag Ultra Derivatization Kit (Waters Corporation) according to the manufacturer’s instructions. An aliquot (50 µL) of the derivatized AA extract was transferred to UPLC vials. AAs were measured as described previously (54).

Fluorometric Starch Quantification.

Total starch content was measured from rosette leaves of 5-wk-old A. thaliana WT, and bZIP triple (bzip2/-11/-44) or quintuple (bzip1/-2/-11/-44/-53) mutants grown under long-day conditions at 100 µmol photons m⁻² s⁻¹ and 60% relative humidity using an Amplex Red–based fluorescent quantification assay. A detailed protocol can be found in SI Appendix.

RNAseq.

To obtain a global transcript profile of the WT, bzip2/-11/-44, and bzip1/-2/-11/-44/-53, we cultivated plants at 100 µmol m⁻² s⁻¹ under long-day conditions until the onset of flowering of the primary inflorescence. At this developmental stage, we harvested at ZT0 flowers, mature and young leaves, as well as an axillary branch enriched tissue (rosette core) from three individual plants per genotype and pooled the material to create each biological replicate. Total RNA was extracted from three biological replicates per genotype and tissue. DNA libraries were prepared from 500 ng of total RNA with oligo-dT capture beads for poly-A-mRNA enrichment using the TruSeq Stranded mRNA Library Preparation Kit (Illumina; https://www.illumina.com) according to manufacturer’s instructions (½ volume). Reads of each sample were mapped to the Arabidopsis genome (Col-0 TAIR10 genome release) as reference. Genes with an adjusted P-value ≤ 0.05 (method: P-adjust “BH” correction) and an absolute log₂FC ≤ −1 (down-regulated) or log₂FC ≥ 1 (up-regulated) were considered as differentially expressed (Dataset S1). Raw and processed data from RNAseq of this article can be found in Gene Expression Omnibus (GSE252502). A detailed description can be found in SI Appendix.

Quantitative Real-Time PCR (qRT-PCR).

For transcript analysis total RNA was extracted from plant tissue using the Trizol extraction method (55). RNA was converted to complementary DNA using H⁻ reverse transcriptase and a mix of random nonamer and oligo dT primers. Quantification of transcript abundance was done by qRT-PCR using SYBR green-based detection. Data were obtained from three biological replicates each derived from plant material of three individual plants. Presented are mean expression values normalized to UBIQUITIN5 expression. Gene-specific oligonucleotide primers used for qRT-PCR analyses are summarized in SI Appendix, Table S2.

ATAC-Sequencing.

A detailed protocol of ATAC-sequencing of protoplasts of WT and Pro35S::bZIP11-GR plants treated with solvent control, DEX, cycloheximide, or both (DEX+cycloheximide) is presented in SI Appendix.

DAPseq.

DAPseq data were obtained online from the Anno-J Networked Genome Browser (http://neomorph.salk.edu/dap_family.php?active=DAP+data/bZIP_tnt). A detailed description of the DAP-seq experiment was published previously (36).

PTAs.

Protoplast transfection assays were performed as previously described (56) with minor modifications (21). β-glucuronidase (GUS) enzyme assays were performed after 16 h incubation. Construction of effector plasmids was described previously (23). SWEET15 and GAT1_2.1 reporter constructs were generated in this study, by PCR-amplifying 1.6 kb of the SWEET15 and 1 kb of the GAT1_2.1 promoter using sequence-specific primers that introduced NcoI and PstI restriction sites into the PCR product. After restriction of the PCR products with NcoI and PstI, the resulting promoter fragments were ligated into the multiple cloning site of the pBT10-GUS vector, upstream of the GUS reporter gene. For mutagenesis of putative bZIP binding sites within the SWEET15 and GAT1_2.1 reporter constructs, the QuickChange II site-directed mutagenesis kit (Agilent Technologies) was used, in combination with the mutagenic primers given in SI Appendix, Table S2.

C and N Measurements.

We determined C and N contents from rosettes and flowers of WT and triple mutants prior axillary branching. Plants were grown on soil under long-day conditions (16 h light/8 h darkness) at 21 °C, an irradiance of 100 µmol photons m⁻² s⁻¹, and a relative humidity of 60%. C and N content measurements were carried out using a UNICUBE elemental analyzer (Elementar GmbH, Germany). A detailed description can be found in SI Appendix.

Gas Exchange Measurements.

We measured the CO₂ assimilation rate according to published protocols (57). In brief, we performed gas exchange measurements from detached leaves of 5-wk-old WT and triple mutant plants grown on soil under long-day conditions (16-h light/8-h darkness) at 21 °C, an irradiance of 120 µmol photons m⁻² s⁻¹, and a relative humidity of 60%. Leaves were cut under water to avoid xylem embolism, and the petiole was kept in deionized water throughout the measurement period. CO₂ uptake was recorded using a custom-made setup with two parallel water-cooled cuvettes with a gas stream of 1 L min⁻¹. The gas composition was adjusted and controlled by mass flowmeters and adjusted to 52.5% RH at 20 °C and 400 ppm CO₂. Leaves were illuminated by two LEDs at 120 µmol photons m⁻² s⁻¹. Recordings were performed with two infrared gas analyzers (LI 7000; Li-Cor, Lincoln, NE, USA).

Statistics.

Statistically significant differences between genotypes at individual timepoints are presented either in the figures or for reasons of clarity given in Dataset S2. Normality of data was checked using the Shapiro–Wilk test. For pairwise comparisons either a two-tailed Student’s t-test (for normal distributed data) or the Mann-Whitney U test (for not-normal distributed data) was applied (n.s., not significant; *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001). For multiple comparisons either the ANOVA test (Tukey post hoc) (for normal distributed data) or Kruskal–Wallis ANOVA test (for not-normal distributed data) was used.

Supplementary Material

Appendix 01 (PDF)

Dataset S01 (XLSX)

Dataset S02 (XLSX)

Data, Materials, and Software Availability

RNAseq datasets have been deposited in Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE252502) (58). All other data are included in the manuscript and/or supporting information.