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Keywords:

  • INDETERMINATE DOMAIN;
  • flowering;
  • sugar metabolism;
  • sucrose synthase;
  • sucrose transporter;
  • Arabidopsis

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

There has been a long-standing interest in the role played by sugars in flowering. Of particular interest is how sugar-related signals are integrated into flowering genetic pathways. Here, we demonstrate that the INDETERMINATE DOMAIN transcription factor AtIDD8 regulates photoperiodic flowering by modulating sugar transport and metabolism. We found that whereas AtIDD8-deficient idd8 mutants exhibit delayed flowering under long days, AtIDD8-overexpressing plants (35S:IDD8) show early flowering. In addition, the sucrose synthase genes SUS1 and SUS4 were upregulated in 35S:IDD8 plants but downregulated in idd8 mutants, in which endogenous sugar levels were altered. AtIDD8 activates the SUS4 gene by binding directly to its promoter, resulting in promoted flowering in SUS4-overexpressing plants. SUS4 expression also responds to photoperiodic signals. Notably, the AtIDD8 gene is suppressed by sugar deprivation. Therefore, we conclude that AtIDD8 regulation of sugar transport and metabolism is linked to photoperiodic flowering.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The timing of flowering is coordinately regulated by diverse internal and external factors. External factors that affect flowering include changes in day length, or photoperiod, and exposure to low temperatures for a long period (Levy and Dean, 1998; Corbesier and Coupland, 2006; Kim et al., 2009). Ambient temperature also influences the floral transition (Blázquez et al., 2003; Kumar and Wigge, 2010). These external signals interact with internal cues through an intricate signaling network to synchronize plant developmental status with changing environmental conditions (Mouradov et al., 2002).

A fundamental but poorly understood question relates to the physiological or metabolic control of flowering. It has been reported that sucrose promotes flowering in many plant species (Bernier et al., 1993; Corbesier et al., 1998; Gibson, 2005; Corbesier and Coupland, 2006). Sucrose levels increase in the phloem sap and apical buds upon exposure of plants to flowering-inductive stimuli (Corbesier et al., 1998). Application of sucrose to the shoot apical meristem (SAM) also promotes flowering (Bagnall and King, 2001). However, the effects of sucrose vary depending on the genetic background of the plant and the growth conditions (Ohto et al., 2001; Gibson, 2005). For example, sucrose promotes flowering when fed to plants at low concentrations, but delays flowering when applied at high concentrations (Ohto et al., 2001). Uncertainties regarding the role of sucrose in floral transition reflect the complexity of genetic and molecular mechanisms linking the timing of flowering with sugar metabolism.

Endogenous sugar levels are intimately associated with photosynthetic carbon assimilation (Moore et al., 2003; Smith et al., 2005). Consequently, photosynthetic activities influence flowering. High irradiance-mediated photosynthesis promotes flowering in Anagallis arvensis (Bernier et al., 1993). In contrast, elimination of atmospheric CO2 significantly delays flowering under long days (LDs) (Kinet et al., 1973). In Arabidopsis, it has been shown that photosynthesis in leaves exposed to high irradiance under red light-rich LDs accelerates flowering via the FLOWERING LOCUS T (FT)-dependent pathway (King et al., 2008). Under identical light conditions, endogenous sucrose levels are elevated in the leaves and shoot apex, and photosynthetically regulated genes, such as SUCROSE-PROTON SYMPORTER 2 (SUC2) and SUCROSE SYNTHASE 1 (SUS1), are induced, supporting the connection between photosynthesis and flowering (Kinet et al., 1973; King et al., 2008).

More direct evidence supporting the association of flowering with sugars has been inferred from studies of sugar contents in late-flowering mutants of Arabidopsis. It has been observed that excess starch accumulates in the leaves of the late-flowering GIGANTEA (GI)-deficient mutant (Eimert et al., 1995). The FT gene is also implicated in sugar regulation of photoperiodic flowering. Whereas the late-flowering phenotype of the CONSTANS (CO)-defective mutant is partly restored by sucrose feeding in darkness, the ft mutant phenotype is insensitive to sucrose both in light and darkness (Roldán et al., 1999; Ohto et al., 2001), suggesting that sucrose-mediated signals are incorporated into the photoperiod flowering pathway, probably downstream of CO but upstream of FT.

Recent studies of INDETERMINATE DOMAIN (IDD) transcription factors have shed light on the molecular link between sugars and flowering. Some IDD members reportedly function in sugar metabolism and floral transition. The maize ID1 protein functions as a transcriptional regulator of floral transition (Colasanti et al., 1998). Notably, in the late-flowering id1 mutant, genes involved in photosynthesis and carbon fixation are induced, suggesting that floral transition and carbon assimilation are interconnected (Coneva et al., 2007). ID1 gene homologs have been identified in rice and Sorghum (Colasanti et al., 1998; Matsubara et al., 2008; Park et al., 2008). The Arabidopsis genome contains sixteen IDD proteins, collectively designated AtIDDs (Colasanti et al., 2006). We therefore hypothesized that at least some AtIDD members may be related to sugar metabolism, and thereby might also be involved in the control of the floral transition.

In this work, we demonstrate that sugar metabolism is intimately linked with the timing of flowering, in which the AtIDD8-SUS4 module plays a critical role. Whereas expression of the AtIDD8 gene was unaffected by changes in day length, it was suppressed under sugar deprivation occurring in darkness or caused by impaired photosynthesis. The SUS4 gene was directly regulated by AtIDD8. It was also regulated by photoperiodic signals independent of AtIDD8. Furthermore, endogenous sugar status also influenced the SUS4 gene via a feedback loop. We therefore propose that photosynthetic and photoperiodic signals are integrated via the AtIDD8-SUS4 module to regulate photoperiodic flowering.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

AtIDD8 protein is a transcriptional activator

As an initial step towards understanding the relationship between sugars and flowering, we obtained a series of T-DNA insertional mutants of AtIDD genes available from public databases and examined their flowering phenotypes. Among the mutants examined, only AtIDD8-deficient mutants showed delayed flowering. A knock-out mutant of the AtIDD3 gene, which shows the highest homology with AtIDD8 (Colasanti et al., 2006), exhibited no discernible phenotypes (Figure S1). We therefore chose the AtIDD8 gene for further studies.

Like known IDD proteins, AtIDD8 is characterized by having four zinc fingers (ZFs) (Figures 1a,b), with the first two being C2H2 types and with the last two being C2HC types. One structural distinction between AtIDD8 and maize ID1 is a shorter ZF1–ZF2 spacer (Figure 1b).

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Figure 1.  Transcriptional activation activity of AtIDD8. (a) Sequence comparison of AtIDD8 and related INDETERMINATE DOMAIN (IDD) proteins. Four zinc-finger (ZF) motifs are underlined; •, cysteine residues; ○, histidine residues. (b) Protein structure of AtIDD8. Four ZF motifs are present in the N-terminal region. Black bars indicate the ZF motifs. (c) Subcellular localization of AtIDD8. The GFP-AtIDD8 fusion gene was transiently expressed in onion epidermal cells. (d) Reporter and effector vectors used for transient expression assays in Arabidopsis protoplasts. GAL4 transient expression assays were carried out as described previously (Miura et al., 2007). The Renilla luciferase gene served as an internal control to normalize values in individual assays. (e) AtIDD8 constructs assayed and relative GUS activity assays. Five independent measurements were averaged. Bars indicate the standard error of the mean. ARF5M is a positive control (Tiwari et al., 2003).

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To determine the subcellular localization of AtIDD8, a green fluorescence protein (GFP)-IDD8 fusion construct, in which the GFP-coding sequence was fused in-frame to the 5′ end of the AtIDD8 gene, was transiently expressed in onion epidermal cells. The GFP signals in those cells were detected exclusively in the nucleus (Figure 1c).

We next examined whether AtIDD8 is a transcriptional regulator using a GAL4 transient expression system in Arabidopsis protoplasts (Miura et al., 2007). A full-size AtIDD8 sequence was fused in-frame to the 3′ end of the GAL4 DNA-binding domain-coding sequence in the effector vector (Figure 1d). This vector, a construct containing the GUS (β-glucuronidase) reporter gene and a vector containing the Renilla luciferase gene, were co-transformed into Arabidopsis protoplasts (Figure 1d). Expression of the AtIDD8 sequence elevated GUS activity by approximately fivefold (Figure 1e, right panel), demonstrating that AtIDD8 protein is a transcriptional activator.

To map a potential transactivation domain, several constructs expressing truncated AtIDD8 sequences were also assayed (Figure 1e, left panel). Whereas expression of the ΔC sequence exhibited no discernible effect, that of the ΔN sequence elevated GUS activity by more than threefold (Figure 1e, right panel). In particular, the expression of the ΔN–N sequence, consisting of residues 171–320, elevated GUS activity by approximately sevenfold. In contrast, expression of the ΔN–C sequence, which included residues 321–466, elevated GUS activity by only twofold. Collectively, these findings indicate that the transactivation domain coincides with the ΔN–N sequence.

Quantitative real-time RT-PCR (qRT-PCR) analyses revealed that AtIDD8 is expressed throughout the life of the plant, but at relatively higher levels in young seedlings (Figure S2a). In addition, whereas transcript levels were relatively high in vegetative organs, they were lower in flowers and siliques (Figure S2b). These expression patterns differ from those of ID1, which is expressed primarily in immature leaves (Colasanti et al., 1998). Together with the structural difference between these two proteins (Figure 1b), these observations suggest that AtIDD8 and ID1 are functionally distinct in some aspects, although both play a role in the floral transition.

idd8 mutants exhibit delayed flowering

Three independent knock-out mutants with T-DNA insertions at different loci within the AtIDD8 gene were obtained from the mutant pool deposited into the Arabidopsis Biological Resource Center (Figure 2a). The absence of gene expression was verified by RT-PCR in each case. All three mutants exhibited late flowering to a similar degree when grown under LDs (Figure 2b), as determined by counting the number of leaves at flowering (Figure 2c) and by the number of days to bolting (Figure 2d). Whereas control plants (Col-0) flowered at a total leaf number of approximately 12, idd8 mutants initiated flowering at a total leaf number of 18–22 under our growth conditions (Figure 2c).

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Figure 2.  Delayed flowering in the idd8 mutants. (a) Mapping of T-DNA insertion sites in the idd8 mutants. (b) Flowering phenotype of the idd8 mutants. Five-week-old plants grown in soil under long days (LDs) were photographed. (c and d) Determination of leaf numbers in individual developmental phases (c) and of days to bolting (d). Approximately 30 plants grown under LDs were counted and averaged in each assay. Bars indicate the standard error of the mean. Statistical significance was determined by a Student’s t-test (*P < 0.01). (e) Expression of FT, AP1, SOC1 and FLC in idd mutants. Aerial parts of 2-week-old plants grown on MS-agar plates harvested at ZT16 were used for total RNA extraction. Transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate the standard errors of the means. Statistical significance was determined by a Student’s t-test (*P < 0.01).

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Notably, counting leaves at different leaf developmental stages revealed that, whereas the juvenile-to-adult phase transition was unaffected, the vegetative-to-reproductive phase transition (floral transition) was significantly delayed in the idd8 mutants (Figure 2c). These observations indicate that AtIDD8 regulates flowering primarily by modulating the reproductive phase change. This is distinct from the role of ID1, which affects both vegetative and reproductive phase changes in maize (Colasanti et al., 1998). As all three idd8 mutants exhibited similar flowering phenotypes, the idd8-1 mutant was used for subsequent analyses.

The expression of flowering time genes was examined by qRT-PCR in the idd8-1 mutant. We found that FT expression was reduced by approximately 50% in the mutant (Figures 2e and S3). FT transcript levels in the idd8-1 idd3-1 double mutant were similar to those seen in the idd8-1 mutant, confirming that AtIDD3 does not contribute to flowering. APETALA1 (AP1), which functions downstream of FT, was also suppressed in the idd8-1 mutant. The SOC1 transcript level was also reduced in the mutant, but to a lesser degree than the reduction in FT and AP1 transcripts. In contrast, FLC expression was not significantly affected in the mutant. CO and GI expression was also not altered in the mutant (Figure S3). These observations indicate that AtIDD8 exerts its role primarily by modulating FT. Accordingly, the idd8-1 mutant overexpressing FT (idd8-1X35S:FT) exhibited early flowering comparable to that in the 35S:FT transgenic plants (Figure S4).

The senescing process is accelerated in 35S: IDD8 transgenic plants

To further explore the role of AtIDD8 in plant developmental timing, we produced Arabidopsis transgenic plants overexpressing AtIDD8 under the control of the Cauliflower mosaic virus (CaMV) 35S promoter. At least 35 independent, single-insertional homozygotic 35S:IDD8 transgenic plants were obtained by herbicide selection and analysis of segregation ratios.

35S:IDD8 transgenic plants exhibited early flowering when grown in soil (see below); however, they exhibited more complex phenotypes when grown on half-strength MS-agar plates devoid of sugar (MS-agar plates, hereafter). Whereas approximately half of the transgenic lines, designated 35S:IDD8s, exhibited severe phenotypic changes, the other half, designated 35S:IDD8m, were phenotypically similar to control seedlings (Figure 3a). 35S:IDD8s seedlings exhibited reduced growth with pale-green leaves, and they eventually underwent senescence without producing flowers. Consistently, whereas the chlorophyll content of 35S:IDD8m seedlings was not significantly altered compared with controls, that of the 35S:IDD8s seedlings was reduced by more than 70% (Figure 3b). These observations were confirmed by analysis of senescence-related genes. Senescence-associated gene 12 (SAG12) and SENESCENCE 4 (SEN4) expression was significantly elevated in 35S:IDD8s seedlings (Figure S5). Analysis of AtIDD8 transcript levels indicated that the phenotypic differences observed between 35S:IDD8m and 35S:IDD8s seedlings were caused by gene dosage effects: AtIDD8 transcript levels were significantly higher in the latter than in the former (Figure 3c).

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Figure 3.  Disrupted chloroplast development in 35S:IDD8 transgenic plants. (a) Seedling phenotypes. Shown are 3-week-old seedlings grown on MS-agar plates. 35S:IDD8 transgenic plants exhibit two phenotypes: 35S:IDD8s (severe) and 35S:IDD8m (mild). (b) Measurement of chlorophyll content. Leaves of plants shown in (a) were used for chlorophyll extraction. Five measurements were averaged. Bars indicate standard errors of the means. (c) AtIDD8 expression. Aerial parts of the plants shown in (a) were used for total RNA extraction, and transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate the standard errors of the means. (d) Transmission electron microscope (TEM) images of chloroplasts. The leaves of 2-week-old plants grown on MS-agar plates were subject to TEM: Ch, chloroplast; Gr, grana; Pl, plastoglobule (lipid body); St, starch grain. Scale bars: 1 μm (left panels); 0.1 μm (right panels).

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Reduced chlorophyll content observed in the 35S:IDD8s leaves suggested that chloroplast development could be perturbed. Electron microscopic analysis revealed that the overall architecture of chloroplasts was severely disrupted in the 35S:IDD8s leaves. Grana stacks were poorly developed, and plastoglobules and starch grains were smaller than those observed in control leaves (Figure 3d). In addition, genes reflecting chloroplast development were also significantly suppressed in 35S:IDD8s transgenic plants (Figure S5).

35S:IDD8s phenotypes are rescued by glucose

35S:IDD8s phenotypes are similar to those observed in Arabidopsis mutants with defective sugar metabolism or transport (Gottwald et al., 2000), suggesting that AtIDD8 functions in these activities. Therefore, we asked whether administration of various sugars could rescue 35S:IDD8s phenotypes.

Seeds of 35S:IDD8 transgenic plants and also the idd8-1 mutant were germinated and grown on MS media supplemented with sugars. Whereas feeding with 1% glucose efficiently rescued 35S:IDD8s phenotypes (Figure 4a), feeding with 2% sucrose had only a marginal effect (Figure 4b). At 6% sucrose, we observed dwarfed growth of both control and 35S:IDD8 seedlings, which was probably the result of osmotic stress. Treatment with 2% mannitol simply imposed osmotic stress on seedlings without rescuing phenotypic defects (Figure S6), indicating that the effects of glucose on 35S:IDD8s phenotypes are metabolic rather than osmotic.

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Figure 4.  Effects of the administration of sugar on 35S:IDD8s phenotypes. (a) Effects of glucose. Seeds were germinated and grown on MS-agar plates supplemented with 1% glucose. Two-week-old seedlings were photographed. (b) Effects of sucrose. Sucrose feeding was carried out as in (a). Three-week-old seedlings were photographed. (c) Measurements of endogenous starch content. Two-week-old whole plants grown on MS-agar plates were used for starch extraction. The value 1 is equal to 0.3 mg g−1 fresh weight. Five measurements were averaged; ZT, zeitgeber time. (d) Measurements of endogenous sugar content. Shoots and roots of 2-week-old plants grown on MS-agar plates harvested at ZT16 were used for sugar extraction. The value 1 is equal to 0.9 and 0.3 mg g−1 of fresh weight for glucose and sucrose, respectively. Five measurements were averaged. Bars indicate standard errors of the means.

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It is known that the endogenous level of starch is reduced when sucrose metabolism is interrupted (Bieniawska et al., 2007). We observed that there were fewer starch grains of smaller size in the 35S:IDD8s chloroplasts (Figure 3d), suggesting that starch content is reduced in the 35S:IDD8s seedlings. To verify this notion, plants were grown on MS-agar plates for 2 weeks under LDs, and whole plants were harvested at different zeitgeber time (ZT) points. Starch content was low when transgenic and mutant seedlings were harvested at ZT0, as observed in control seedlings (Figure 4c). However, the starch content gradually increased with time after ZT0, peaking at ZT16 in control plants. Notably, the rate of increase was much less in 35S:IDD8s seedlings, probably because of malfunctioning chloroplasts. Similar patterns in starch accumulation were observed in 35S:IDD8m seedlings, but to a lesser degree (data not shown). In contrast, the rate of starch increase was greater in idd8-1 seedlings. These observations indicate that sugar metabolism is altered in transgenic and mutant seedlings. Starch levels were also higher in a few late-flowering mutants, such as gi-2 and fca-9, but were unaltered in ft-10, soc1-101 and fve-3 mutants (Figure S7), suggesting that the physiological basis of late-flowering phenotypes is not the same in different late-flowering mutants. Alternatively, starch accumulation may not be linked directly with flowering phenotypes, at least in Arabidopsis.

We next measured endogenous levels of sucrose and glucose in shoots and roots of 2-week-old 35S:IDD8s and idd8-1 seedlings grown on MS-agar plates and harvested at ZT16. Overall sucrose and glucose levels were relatively higher in the shoots and roots of idd8-1 seedlings compared with control and 35S:IDD8s seedlings (Figure 4d), similar to elevated levels of endogenous starch observed in idd8-1 seedlings (Figure 4c). In 35S:IDD8s shoots, sucrose and glucose levels were similar to those seen in control shoots. In contrast, sucrose levels were much lower in 35S:IDD8s roots than in control roots, whereas glucose levels did not significantly differ in root samples. These observations support the hypothesis that AtIDD8 plays a role in sucrose transport as well as in sugar metabolism.

AtIDD8 binds to a consensus motif in the SUS4 promoter

Whereas 1% glucose efficiently rescued 35S:IDD8s phenotypes, 2% sucrose did not (Figure 4a,b). In addition, sugar levels were altered in 35S:IDD8s and idd8-1 seedlings (Figure 4c,d), showing that AtIDD8 functions in sucrose transport and metabolism.

To analyze this notion mechanistically, we carried out qRT-PCR analysis of genes encoding sucrose transporters (SUC1SUC9) and sucrose synthases (SUS1SUS6) in transgenic and mutant plants. As 35S:IDD8s transgenic plants senesced prematurely, 35S:IDD8m transgenic plants were used for subsequent analyses. Among the SUC genes examined, expression of SUC2, SUC6, SUC7 and SUC8 genes was affected in both 35S:IDD8m transgenic and idd8-1 mutant plants. Whereas it was repressed in the transgenic plants, it was slightly upregulated in the idd8-1 mutant (Figure 5a), indicating that AtIDD8 negatively regulates SUC expression.

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Figure 5.  AtIDD8 activation of SUS4. In (a) and (b), transcript levels were determined by qRT-PCR. Biological triplicate samples were averaged. Bars indicate standard errors of the means. (a) SUC expression as determined by qRT-PCR in 35S:IDD8m and idd8-1. (b) SUS expression as determined by qRT-PCR in 35S:IDD8m and idd8-1. The y-axis represents a logarithmic scale to better compare fold changes. (c) Binding of AtIDD8 to a conserved sequence in the SUS4 promoter. Binding activities were examined by EMSA. A consensus ID1 binding sequence (Kozaki et al., 2004) was included as a control. The core sequence is underlined. Recombinant AtIDD8 protein expressed in Escherichia coli cells and end-labeled DNA fragments were used. The (–) lane represents a control without the addition of AtIDD8 protein. Varying quantities of unlabeled DNA fragments were added as competitors. (d) GUS activity assays in Arabidopsis protoplasts. The (–) indicates the reporter vector without the SUS4 promoter sequences. Five measurements were averaged (lower panel). Bars denote standard errors of the means. Vector, control vector without the AtIDD8 gene. (e) Early flowering of 35S:SUS4 transgenic plants. Four-week-old plants grown in soil under long days (LDs) were photographed (upper panel). Two representative transgenic lines (T11 and T14), each consisting of 20 plants, were counted (lower panel). Statistical significance was determined by a Student’s t-test (*P < 0.01). (f) Early flowering of the idd8-1 mutant overexpressing the SUS4 gene. The counting of rosette leaves and determination of statistical significance were carried out as described in (e).

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Transcript levels of SUS1 and SUS4 genes were also greatly altered in both 35S:IDD8m transgenic and idd8-1 mutant plants. Specifically, SUS1 transcript levels were elevated by approximately fivefold in 35S:IDD8m seedlings and were slightly reduced in the idd8-1 mutant (Figure 5b). Notably, SUS4 transcript levels were robustly elevated by more than 14-fold in 35S:IDD8m seedlings and were reduced by approximately 30% in the idd8-1 mutant. The At1g22650 gene, which encodes a putative invertase, also showed an expression pattern similar to those of SUS1 and SUS4 genes, but with a lesser degree of alterations: its transcript levels were elevated in 35S:IDD8m seedlings but were reduced in the idd8-1 mutant (Figure 5b). These observations indicate that AtIDD8 regulates sugar metabolism by modulating SUS1 and SUS4, and support the 35S:IDD8s seedling phenotypes, which were rescued efficiently by glucose but not by sucrose.

Given that AtIDD8 is a transcriptional activator (Figure 1e), we asked whether it directly regulates SUS genes. Sequence analysis of SUS gene promoters revealed that the SUS4 promoter contains a conserved CTTTTGTCC motif (Figure 5c), which is similar to the binding sequence (CTTTCTCTTT) of the DNA binding with one finger (DOF) transcription factors possessing a single ZF motif (Schneidereit et al., 2008). Furthermore, it has been shown that the maize ID1 protein binds to a contiguous 11-bp TTTGTCGTTTT sequence (Kozaki et al., 2004).

To determine whether AtIDD8 binds to the conserved sequence motif within the SUS4 gene promoter (SUS4-BS), we produced recombinant AtIDD8 protein containing the N-terminal IDD domain (residues 1–330) as a maltose binding protein (MBP) fusion in Escherichia coli cells, and carried out gel mobility shift assays using an end-radiolabeled SUS4-BS DNA fragment. We found that AtIDD8 protein bound to the SUS4-BS sequence (Figure 5c). In addition, binding of AtIDD8 to the SUS4-BS sequence was significantly decreased in the presence of unlabeled SUS4-BS, but was reduced to a lesser degree by competition with an unlabeled mutant form (SUS4-mBS) of the binding site. In contrast, AtIDD8 protein did not bind to the ID1-BS sequence (data not shown), indicating that AtIDD8 binding is specific to the SUS4 gene promoter, and further supporting a functional distinction between ID1 and AtIDD8. MBP protein alone did not bind to the SUS4-BS sequence (data not shown).

To examine the binding of AtIDD8 to the SUS4 promoter in planta, we carried out transient co-expression assays in Arabidopsis protoplasts using a series of GUS reporter constructs, in which three copies of the SUS4-P or the mSUS4-P sequence were subcloned upstream of the 35S minimal promoter fused to a GUS gene and a p35S:IDD8 effector plasmid (Figure 5d, upper panel). When the pSUS4-P reporter construct was co-expressed with the effector vector, GUS activity was elevated by more than threefold relative to the control reporter construct (pMin35S) (Figure 5d, lower panel). In contrast, co-expression with the pmSUS4-P reporter construct did not influence GUS activity, showing that AtIDD8 regulates the SUS4 gene.

As AtIDD8 directly regulates SUS4, we predicted that SUS4 upregulation would promote flowering. As expected, transgenic plants overexpressing SUS4 (35S:SUS4) exhibited early flowering under both LDs and SDs (Figures 5e, S8 and S9), and expression of FT and SOC1 is also accordingly altered (Figure S10). Furthermore, SUS4 overexpression in the idd8-1 mutant efficiently rescued the late-flowering phenotype (Figure 5F), confirming the positive regulation of SUS4 by AtIDD8.

AtIDD8-SUS4 signals regulate photoperiodic flowering

When grown in soil under LDs, 35S:IDD8 transgenic plants were largely indistinguishable from control plants in vegetative growth. However, these transgenic plants flowered earlier than the control plants (Figure 6a). 35S:IDD8 transgenic plants also showed early flowering under SDs, particularly when assayed by days to bolting. In contrast, although the idd8-1 mutant was late flowering under LDs, it did not exhibit late flowering under SDs. These observations indicate that AtIDD8 is associated with photoperiodic flowering.

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Figure 6.  Photoperiodic regulation of SUS4. In (b–d), transcript levels were determined by qRT-PCR. Biological triplicate samples were averaged. Bars indicate standard errors of the means. (a) Flowering phenotypes under different light regimes. Plants were grown in soil either under long days (LDs) or short days (SDs) until flowering. Thirty plants were counted and averaged for each plant group. Values represent means ± standard errors. Statistical significance was determined by a Student’s t-test (P < 0.05). (b) Effects of daylength on SUS4 gene expression. In the graph, x-axis numbers indicate zeitgeber time (ZT) points. Plants were germinated and grown on MS-agar plates for 2 weeks, either under LDs or SDs. Whole plants were harvested at the indicated time points for total RNA extraction. (c) SUS4 gene expression in idd8-1. Two-week-old plants grown on MS-agar plates either under LDs or SDs were used for total RNA extraction. (d) SUS4 gene expression in photoreceptor mutants. Two-week-old plants grown on MS-agar plates were used for RNA extraction.

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Interestingly, measurements of AtIDD8 and SUS4 transcript levels in control plants grown either under LDs or SDs revealed that AtIDD8 transcript levels were unaffected by day-length changes (Figure 6b). By contrast, SUS4 was upregulated in plants grown under LDs, but its transcript levels were significantly reduced in plants grown under SDs, indicating that SUS4 rather than AtIDD8 is responsive to photoperiod. Notably, SUS4 expression was still influenced by daylength changes in an idd8-1 background (Figure 6c), indicating that the effects of daylength changes on the SUS4 gene are independent of AtIDD8.

In agreement with the photoperiodic regulation of SUS4, SUS4 transcript levels were reduced in phya and phyb mutants, which have mutations in genes encoding phytochrome-A and -B photoreceptors, respectively (Figure 6d). Together, these observations illustrate that the SUS4 gene is related to photoperiod flowering. SUS1 also showed an expression pattern similar to that of SUS4 in phy mutants, and under different light regimes (Figure S11).

AtIDD8 is suppressed under sugar deprivation

Finally, we determined the nature of input signals regulating the AtIDD8 gene. Extensive gene expression analysis under various growth conditions revealed that AtIDD8 transcript levels were markedly reduced in plants incubated in darkness for longer than 24 h (Figure 7a). SUS4 transcript levels were also reduced, but to a lesser degree, under identical conditions. The fact that shorter incubation times in darkness did not have a significant effect on AtIDD8 suggests that the dark effect was metabolic rather than related to a light/dark response.

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Figure 7.  Regulation of AtIDD8 by sugar deprivation, and working model of sugar-mediated photoperiodic flowering. (a) Dark suppression of AtIDD8. Two-week-old plants grown on MS-agar plates were incubated either in light or in darkness for up to 3 days (d), and AtIDD8 transcript levels were determined by qRT-PCR. Biological triplicate samples were averaged. Bars indicate standard errors of the means. Statistical significance was determined by a Student’s t-test (*P < 0.01). (b) Effects of glucose on AtIDD8 gene expression in darkness. Two-week-old plants grown on MS-agar plates were incubated in darkness for 2 days in the presence of 1% glucose. AtIDD8 transcript levels were determined as described in (a). (c) Effects of a photosynthesis inhibitor on AtIDD8 expression. Two-week-old plants grown on MS-agar plates were incubated in light in the presence of 100 μM 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU). AtIDD8 transcript levels were determined as described in (a). (d) Working model of the roles of AtIDD8 and SUS4 in photoperiodic flowering. SUS4-mediated sugar metabolism constitutes a photoperiod flowering pathway. The AtIDD8 gene integrates signals of endogenous sugar levels into the SUS4 pathway. The SUS4 gene is also regulated by endogenous sugar levels, such as glucose, via feedback regulation.

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To test this hypothesis, plants were incubated in darkness for 2 days, either in the presence or absence of 1% glucose, and AtIDD8 transcript levels were determined by qRT-PCR. Reduced AtIDD8 transcripts were restored to a level comparable with that seen in light-grown plants in the presence of glucose (Figure 7b), supporting the metabolic nature of dark effects on AtIDD8 gene expression. Furthermore, when plants were grown in light in the presence of 100 μm 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), which specifically inhibits photosynthesis (Metz et al., 1986), AtIDD8 expression was suppressed, as observed in dark-grown plants (Figure 7c). Therefore, we concluded that AtIDD8 expression is suppressed under sugar deprivation, which occurs in darkness or under impaired photosynthesis.

Overall, our data indicate that AtIDD8 is responsive to levels of endogenous sugars, and integrates those signals into the SUS4-mediated photoperiod flowering (Figure 7d). SUS4 expression is also influenced by endogenous sugar content via feedback regulation, as we observed that SUS4 is induced by feeding with glucose or sucrose (Figure S12). We therefore propose that the AtIDD8-SUS4 module balances metabolic status and photoperiodic flowering, and ensures that floral transition occurs when endogenous sugar levels are sufficient to sustain reproductive success.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

INDETERMINATE DOMAIN proteins are a family of plant-specific, IDD-containing transcription factors (Colasanti et al., 2006). Since the discovery of the maize ID1 protein (Colasanti et al., 1998), a few IDD proteins have been functionally characterized in rice and Arabidopsis. For example, it has been shown that a rice ID1 homolog functions in the initiation of flowering (Matsubara et al., 2008; Park et al., 2008). The Arabidopsis shoot gravitropism 5 (SGR5) gene encodes AtIDD15 protein, which functions in sugar-mediated gravitropic responses (Morita et al., 2006; Tanimoto et al., 2008). However, it is currently unknown how IDD transcription factors regulate downstream signaling governing developmental timing and physiological processes.

AtIDD8/NUTCRACKER (NUC) and its close homologs AtIDD3/MAGPIE (MGP) and AtIDD10/JACKDAW (JKD) have been reported as regulators of root development. Large-scale meta-analysis of several microarray data has shown that AtIDD8/NUC and AtIDD3/MGP are direct targets of SHORT-ROOT (SHR) transcription factor, which plays key roles in specifying the root stem cell niche and radial root patterning (Levesque et al., 2006). Furthermore, AtIDD3/MGP and AtIDD10/JKD regulate root tissue boundaries and asymmetric cell division in roots by modulating SHR and SCARECROW (SCR) activities through a feed-forward loop (Welch et al., 2007; Hassan et al., 2010).

Here, we demonstrated that AtIDD8 also plays an important role in photoperiodic flowering. DNA microarray data in the public database show that AtIDD8 is expressed predominantly in roots (https://www.genevestigator.com/gv/index.jsp), thereby suggesting that AtIDD8 may play a role in transporting a certain root-derived signal to stems/leaves to promote floral transition. However, we found that the AtIDD8 gene is also expressed to a high level in stems and leaves (Figure S2). In particular, the AtIDD8 transcription factor directly regulates the SUS4 gene, linking the endogenous metabolic status with the induction of photoperiodic flowering. Consistent with the notion that sugar metabolism is closely associated with the timing of flowering via AtIDD8 regulation of SUS4, both flowering time and endogenous sugar contents were accordingly altered in 35S:SUS4 transgenic plants as well as in 35S:IDD8 transgenic plants and the idd8-1 mutant, supporting the hypothesis that AtIDD8 expressed in the stems and leaves is related to the induction of photoperiodic flowering.

In plants, developmental phase transitions require significant energy input. Control of sugar metabolism or mobilization is necessary to maintain proper energy resources, particularly during reproductive or floral transitions (Koch, 2004; Pourtau et al., 2006; Baena-González et al., 2007; Baena-González and Sheen, 2008). Therefore, it is not surprising that sugar metabolism is connected with the floral transition and the senescing process. Over the last decades, numerous studies have supported the functional significance of sugars in controlling flowering time, although the genes and regulatory mechanisms underlying the metabolic control of flowering have not yet been characterized at the molecular level. Our data provide direct molecular genetic and physiological evidence supporting the link between sugar levels and flowering.

Whereas AtIDD8 expression is unaffected by feeding with sucrose or glucose, it is suppressed by dark-induced sugar deprivation and impaired photosynthesis. These observations indicate that AtIDD8 is sensitive to endogenous levels of sugars, and modulates SUS4 activity to determine the proper timing of phase transitions. When endogenous sugar levels are sufficiently high, AtIDD8 promotes floral transition to achieve reproductive success by inducing SUS4. However, when endogenous sugar levels drop below the threshold required for proper developmental transitions, AtIDD8 is downregulated, and vegetative growth continues until sufficient amounts of photosynthetic products accumulate. We therefore concluded that sugar accumulation is not the cause of the late flowering of the idd8-1 mutant. Instead, it is envisaged that sugars accumulate in the idd8-1 mutant because a lack of AtIDD8 expression is perceived as a signal of low levels of sugars in the mutant, which is also consistent with the positive role of sugars in the floral transition. This signaling scheme may explain the phenotypes seen in 35S:IDD8s transgenic plants grown on MS-agar plates: their perturbed growth and pale-green leaves could be the result of excessive consumption of endogenous sugars by the elevated SUS4 activity at a time when sugar availability is limited.

Notably, AtIDD8-mediated sugar metabolism is linked to the SUS4-mediated photoperiod flowering. It has been shown that photosynthesis-derived sucrose promotes flowering via the FT gene (King et al., 2008). Late-flowering phenotypes of the autonomous pathway mutants, such as fca, fpa and fve, are rescued by exogenous sucrose (Roldán et al., 1999). By contrast, the late flowering phenotype of the ft mutant is largely insensitive to sucrose feeding, further supporting the critical role of FT in sucrose induction of flowering.

We also found that FT gene expression is reduced in the idd8-1 mutant, which exhibits late flowering under LDs but not SDs. However, the AtIDD8 gene is unaffected by photoperiodic signals. Instead, its target SUS4 is induced under LDs but suppressed under SDs. Furthermore, SUS4 is also induced by feeding with sucrose and glucose, indicating that the SUS4 gene incorporates both photoperiodic and metabolic signals into a unique genetic pathway regulating photoperiodic flowering. We therefore conclude that AtIDD8 contributes to photoperiodic flowering indirectly by modulating SUS4 activity (Figure 7d).

Carbon assimilation is closely related with developmental transitions, such as the floral transition. Our data indicate that metabolic status is directly linked to floral transition. Plants have apparently evolved a way of synchronizing endogenous sugar status with photoperiodic flowering. It seems that the SUS4 gene, as well as unidentified AtIDD8 target genes, could constitute a unique, FT-dependent photoperiod flowering pathway that is further modulated through physiological control. Photoperiod is a potential determinant for durations of sugar accumulation (Bagnall and King, 2001), as well as for floral induction, further supporting a linkage between sugar deprivation-mediated AtIDD8 signaling and photoperiodic flowering.

Our data indicate that the AtIDD8-SUS4 module plays a critical role in controlling developmental transitions by monitoring endogenous sugar status (Figure 7d). However, the proposed signaling scheme could be more complex. Whereas 35S:SUS4 transgenic plants showed early flowering (Figure 5e), the flowering of sus1 sus2 sus3 sus4 quadruple mutants is essentially normal (Barratt et al., 2009). The different flowering phenotypes of the idd8-1 mutant and the sus1 sus2 sus3 sus4 quadruple mutant suggest that AtIDD8 also regulates other genes involved in sugar metabolism and transport, in addition to SUS4. Indeed, we found that several sucrose transporter genes are negatively regulated by AtIDD8. In support of this, transgenic plants overexpressing AtIDD8 are phenotypically similar to the suc2 mutant in that both exhibited reduced growth (Gottwald et al., 2000). The coordination of unidentified targets of AtIDD8 for regulating sucrose transport and sucrose metabolism might be the important part in sugar-mediated floral transition.

It is currently unclear how the AtIDD8 modulation of sugar metabolism and transport influences FT expression. Extensive gene expression profiling revealed that a subset of the SUS genes is upregulated, but that genes encoding invertases are unaffected in 35S:IDD8m plants (data not shown). Whereas invertases produce glucose and fructose from sucrose, SUS enzymes generate UDP-glucose and fructose from sucrose. UDP-glucose is perceived as a signaling molecule as well as a carbon source (Böhringer et al., 1995; Lazarowski et al., 2003). Therefore, UDP-glucose signals, which are mediated by AtIDD8, may regulate FT expression. In addition, sucrose metabolism and distribution in plant tissues is regulated at multiple steps by numerous intrinsic and environmental factors to maintain sugar homeostasis under given growth conditions. Therefore, it is likely that there would be extensive signaling crosstalk between the genetic pathways governing sugar metabolism and transport and the flowering genetic pathways. This notion would explain at least part of the difficulties that we experienced in predicting the effects of sugars on flowering initiation.

Experimental Procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Plant materials and growth conditions

All Arabidopsis thaliana lines used were in the Col-0 background. Plants were grown in a controlled culture room at 22°C with a relative humidity of 55% either under LDs (16-h light/8-h dark) or short days (SDs; 8-h light/16-h dark), with white light illumination (120 μmol photons m−2 sec−1) provided by fluorescent FLR40D/A tubes (Osram, http://www.osram.com). The idd8-1 (SALK-110117), idd8-2 (SALK-065143) and idd8-3 (SALK-124222) mutants were isolated from a mutant pool of T-DNA insertion lines deposited into the Arabidopsis Biological Resource Center (ABRC, Ohio State University). Homozygotic lines were isolated by herbicide selection for three or more generations, and by analysis of segregation ratios. Absence of gene expression in the mutants was verified by RT-PCR before use.

To generate transgenic plants overexpressing AtIDD8 or SUS4, full-size cDNAs were subcloned into the binary pB2GW7 vector under the control of the CaMV 35S promoter. Agrobacterium-mediated Arabidopsis transformation was carried out according to a modified floral-dip method (Clough and Bent, 1998).

Analysis of transcript levels

Total RNA was extracted from appropriate plant materials using the RNeasy Plant Mini Kit (Qiagen, http://www.qiagen.com) and pre-treated with RNAse-free DNase to eliminate contaminating genomic DNA.

Transcript levels were analyzed by qRT-PCR carried out in 96-well blocks using the Applied Biosystems 7500 Real-Time PCR System (Applied Biosystems, http://www.appliedbiosystems.com) and the SYBR Green-I master mix in a volume of 25 μl. The PCR primers were designed using primer express. The two-step thermal cycling profile used was 15 sec at 94°C and 1 min at 68°C. An eIF4A gene (At3g13920) was included in the assays as an internal control to normalize variations in cDNA levels. Reactions were performed in biological triplicates using plant samples harvested separately for each run. The comparative ΔΔCT method was used to evaluate relative quantities of each amplified product. The cycle threshold (CT) was automatically determined for each reaction by the system set with default parameters. PCR specificity was determined by melt-curve analysis of amplified products using the standard method installed in the system. The PCR primers used are listed in Table S1.

AtIDD8 subcellular localization

For detection by fluorescence microscopy, the GFP-coding sequence was fused in-frame to the 5′ end of AtIDD8, and the fusion was subcloned into pBA002 for transient expression in onion epidermal cells. After incubation for 24 h at 23°C, cells were subject to bright-field and fluorescence microscopy using an Olympus BX51TRF fluorescence microscope equipped with a BH2-RFL-T3 UV burner and a DP70 charge-coupled-device camera (Olympus, http://www.olympus-global.com).

Counting the leaf number and phenotypic analysis

Plants were grown in soil until flowering under appropriate growth conditions. The presence of abaxial trichomes, the frequency of hydathodes at leaf margins, and the leaf basal angles were used to discriminate between leaves at different developmental phases. Leaf number was counted using approximately 30 plants for each plant group and then averaged.

Transmission electron microscopy (TEM)

Two-week-old plants grown on MS-agar plates were used. TEM was carried out in the National Instrumentation Center for Environmental Management (NICEM) facilities, Seoul National University. Sections were observed using a JEM1010 transmission electron microscope (JEOL, http://www.jeol.com).

Sugar measurement

For each assay, 0.5 g of plant materials was used to measure the content of soluble sugars using the Sucrose/D-Glucose Assay kit or the Starch Assay kit, according to the manufacturer’s instructions (Megazyme, http://www.megazyme.com). After grinding plant material in liquid nitrogen, soluble sugars were extracted two times with 80% ethanol at 70°C, and fractions were collected and then centrifuged at 12 000 g for 10 min. Ethanol was evaporated from the supernatant using a vacuum evaporator, and the solution volume was adjusted to 400 μl by adding triple-distilled water to measure sugar content. The insoluble pellet was used to measure starch content.

Measurements of chlorophyll content

Measurement of chlorophyll content was carried out as described by Oh et al. (1997). The fourth leaves of 3-week-old plants grown on MS-agar plates were used. Chlorophylls were extracted with N,N-dimethylformamide (DMF), and the extracted solution was incubated at 4°C for 2 h in darkness. Chlorophyll content was assayed by measuring absorbance at 652, 665 and 750 nm using a diode array spectrophotometer (WPA Biowave; Biochrom, http://www.biochrom.co.uk).

Electrophoretic mobility shift assays (EMSA)

An AtIDD8 cDNA was subcloned into the pMAL-c2X expression vector (NEB, http://www.neb.com) containing an MBP-coding sequence. The MBP-AtIDD8 fusion protein expressed in E. coli cells was purified using the pMAL™ Protein Fusion and Purification System (NEB).

DNA fragments ranging from 11 to 23 nucleotides in length were synthesized based on SUS4 promoter sequences, and end-labeled with [γ-32P]dATP using T4 polynucleotide kinase (TaKaRa, http://www.takara-bio.com). Labeled probes were incubated for 30 min at 25°C with 0.5 μg of the MBP-AtIDD8 protein in binding buffer (10 mm Tris–HCl, pH 7.6, 50 mm NaCl, 1 mm EDTA, 5 mm DTT, 5% glycerol) supplemented with 100 ng poly(dI-dC) in the presence or absence of competitor DNA fragments. Reaction mixtures were electrophoresed on 6% native PAGE gels. Gels were dried on Whatman 3MM paper and exposed to X-ray film.

Protoplast transfection assays

For the transient expression assays in Arabidopsis protoplasts, several reporter and effector plasmids were constructed. The pMin35S reporter plasmid contains a minimal CaMV 35S promoter (a region from position −56 to position −8, including the TATA box) and the GUS gene. The pSUS4-P reporter plasmid includes a SUS4 promoter sequence upstream of the 35S minimal promoter. To construct the p35S:IDD8 effector plasmid, the AtIDD8 gene was inserted into an expression vector containing the CaMV 35S promoter and a Nos terminator. The reporter and effector plasmids were co-transformed into Arabidopsis protoplasts by a polyethylene glycol (PEG)-mediated transformation method. The GUS activities were measured by the fluorometric method as described previously (Jefferson et al., 1987). A CaMV 35S promoter-luciferase (LUC) construct was also co-transformed as an internal control. The luciferase assay was carried out using the Luciferase Assay System kit (Promega, http://www.promega.com).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We thank Dae-Jin Yun for providing the reporter and effector vectors for transient expression. This work was supported by the Biogreen 21 (20080401034001) and National Research Laboratory programmes, and by grants from the Plant Signaling Network Research Center (2010-0001453), the National Research Foundation of Korea (2007-03415 and 20090087317), and from the Agricultural R & D Promotion Center (309017-5), Korea Ministry for Food, Agriculture, Forestry and Fisheries.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental Procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Figure S1. Flowering phenotypes of the idd mutants. Five-week-old plants grown in soil under LDs were photographed (top panel). Absence of gene expression in the idd mutants was verified by RT-PCR (bottom panel).

Figure S2.AtIDD8 gene expression patterns. Transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate standard error of the mean. (a) AtIDD8 gene expression at different growth stages. Plants were grown in soil under LDs, and the aerial plant parts were harvested at the indicated time points for total RNA extraction. d, days after germination. (b) AtIDD8 gene expression in different plant tissues. Total RNA samples were extracted from each plant tissue separately.

Figure S3. Flowering gene expression in the idd8-1 mutant. Plants were grown on MS-agar plates under LDs for the indicated time periods. Transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate standard error of the mean. DAG, days after germination.

Figure S4. Flowering phenotypes of idd8-1X35S:FT and its parental mutants. Plants were grown under LDs until flowering. Rosette leaf numbers of 30 plants were counted and averaged for each plant group. Bars indicate standard error of the mean.

Figure S5. Expression of senescence- and photosynthesis-related genes in the idd8-1 and 35S:IDD8 plants. Transcript levels were determined by qRT-PCR. An eIF4A gene (At3g13920) was included in the assays as an internal control for normalizing the variations in cDNA amounts used. The aerial plant parts of 2-week-old plants grown on MS-agar plates were used for RNA extraction. Biological triplicates were averaged. Bars indicate standard error of the mean.

Figure S6. Effects of mannitol and glucose on seedling growth. Seeds were germinated and grown on MS-agar plates supplemented either with 2% glucose or 2% mannitol. Two-week-old seedlings were photographed.

Figure S7. Measurement of starch contents in late flowering mutants. Two-week-old, whole plants grown on MS-agar plates and harvested at ZT16 were used for measurements of starch contents. Five measurements were averaged. Bars indicates standard error of the mean. Statistical significance was determined by a student t-test (*P < 0.01). The value 1 is equal to 1.35mg/g fresh weight.

Figure S8. Flowering phenotypes of 35S:SUS4 transgenic plants under short days (SDs). Plants were grown under SDs until flowering. Rosette leaf numbers of 30 plants were counted and averaged for each plant group. Bars indicate standard error of the mean.

Figure S9. Flowering phenotypes of 35S:IDD8 and 35S:SUS4 transgenic plants under long days (LD). Plants were grown in soil under LDs until flowering. Rosette leaf numbers of 30 plants were counted and averaged for each plant group. Bars indicate standard error of the mean.

Figure S10. Transcript levels of flowering genes in the 35S:SUS4 transgenic plants. Transcript levels were determined by qRT-PCR. Two-week-old, whole plants grown on MS-agar plates and harvested at ZT16 were used for total RNA extraction. Biological triplicates were averaged. Bars indicate standard error of the mean.

Figure S11. Expression of SUS1 under different light regimes and in photoreceptor mutants. Transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate standard error of the mean. (a) SUS1 transcript levels under different light regimes. Plants were germinated and grown on MS-agar plates for 2 weeks either under LDs or SDs. Whole plants were harvested at the indicated time points for total RNA extraction. The numbers on the x-axis indicate ZT points. (b) SUS1 transcript levels in photoreceptor mutants. Two-week-old plants grown on MS-agar plates were used for RNA extraction.

Figure S12. Effects of sugars on the AtIDD8 and SUS gene expression. Two-week-old plants grown on MS-agar plates under LDs were transferred to liquid MS cultures supplemented either with glucose (Glc) or sucrose (Suc) and gently shaken for 24 hours before harvesting plant materials for total RNA extraction. Transcript levels were determined by qRT-PCR. Biological triplicates were averaged. Bars indicate standard error of the mean.

Table S1. Primers used in this study.

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