Gynoecia of the Arabidopsis mutant sty1-1 display abnormal style morphology and altered vascular patterning. These phenotypes, which are enhanced in the sty1-1 sty2-1 double mutant, suggest that auxin homeostasis or signalling might be affected by mutations in STY1 and STY2, both members of the SHI gene family. Chemical inhibition of polar auxin transport (PAT) severely affects the apical–basal patterning of the gynoecium, as do mutations in the auxin transport/signalling genes PIN1, PID and ETT. Here we show that the apical–basal patterning of sty1-1 and sty1-1 sty2-1 gynoecia is hypersensitive to reductions in PAT, and that sty1-1 enhances the PAT inhibition-like phenotypes of pin1-5, pid-8 and ett-1 gynoecia. Furthermore, we show that STY1 activates transcription of the flavin monooxygenase-encoding gene THREAD/YUCCA4, involved in auxin biosynthesis, and that changes in expression of STY1 and related genes lead to altered auxin homeostasis. Our results suggest that STY1 and related genes promote normal development of the style and affect apical–basal patterning of the gynoecium through regulation of auxin homeostasis.
The female reproductive organ of the angiosperm flower, the gynoecium, is one of the most complex and multifunctional organs of the plant. The gynoecium of Arabidopsis consists of two congenitally fused carpels and exhibits two main axes of polarity (Bowman et al., 1999). It is divided into four pattern elements along the apical–basal axis: the stigma, the style, the ovary, consisting of two valves separated by the replum, and the gynophore, a short, solid, stem-like structure. The internal–external, or adaxial–abaxial, axis consists of transmitting tract and placentae adaxially, and style, valve epidermis and replum abaxially. The bilaterally symmetrical ovary is divided into two locules by the septum. Valves occupy lateral positions, whereas repla are medially localized (Bowman et al., 1999). Through each valve, a vascular bundle extends from the base of the gynoecium and terminates just below the valve/style junction. Vascular bundles also run through each replum, bifurcate at the valve/style junction and spread out into large vascular fans in the medial plane of the style.
The distribution and signalling of the phytohormone auxin, indole-3-acetic acid (IAA), has been suggested to affect various processes of polarity establishment and pattern formation in plants (Friml, 2003; Friml et al., 2003; Reinhardt et al., 2003). The response to auxin is dependent on the auxin concentration perceived by the individual cell, and auxin levels are therefore under tight homeostatic control (reviewed by Woodward and Bartel, 2005). Polar auxin transport (PAT), mediated by the putative auxin influx and efflux carriers AUX1 and PIN1-4 and 7, respectively, has been shown to be important in regulating the local auxin concentrations necessary for the establishment of embryo polarity, organ positioning and organ development (Benkova et al., 2003; Friml et al., 2002a,b, 2003; Marchant et al., 2002; Reinhardt et al., 2003).
Auxin synthesis has been shown to occur to different extents in the entire plant (Ljung et al., 2001), and members of two gene families, the YUCCA (YUC) family encoding flavin monooxygenases and the cytochrome P450 family (CYP79B2 and CYP79B3), have been proposed to catalyse important steps in the biosynthesis of auxin (Zhao et al., 2001, 2002), and are therefore potential targets for regulation of the auxin biosynthesis rate in planta. Local concentrations of active auxin can also be regulated through oxidization or conjugation and de-conjugation (reviewed by Woodward and Bartel, 2005). Proteins encoded by the auxin-inducible GH3-like genes can catalyse conjugation of auxin to amino acids in vitro (Staswick et al., 2005), and are probably part of a negative feedback mechanism for preservation of auxin homeostasis. Recent studies on argonaute1 mutants and plants expressing a microRNA resistant version of the AUXIN RESPONSE FACTOR (ARF) 17 indicate important roles for GH3-like protein-mediated auxin conjugation in plant development because the phenotypic alterations observed in these plants correlate with altered GH3-like gene expression and changes in IAA content (Mallory et al., 2005; Sorin et al., 2005).
Several lines of evidence suggest a role for auxin in apical–basal patterning of the gynoecium. Treatment with PAT inhibitors leads to severe defects in gynoecium development such as reduction of the valves in the apical–basal direction, concomitant expansion of the gynophore and stylar regions, premature termination of the lateral veins, and a basalized bifurcation of the medial veins (Nemhauser et al., 2000). Several of these effects are observed in a number of mutants involved in auxin metabolism, transport or signalling.
Mutations in the Petunia gene FLOOZY, a homologue of the Arabidopsis YUC gene, probably involved in auxin synthesis, lead to gynoecial defects similar to those induced by reduced PAT (Tobena-Santamaria et al., 2002). Knockout mutations in the Arabidopsis YUC gene show no phenotypic deviations from wild-type, probably due to functional redundancy with several other related genes (Zhao et al., 2001).
The PIN-FORMED1 (PIN1) gene encodes an auxin efflux facilitator protein that becomes polarly localized in auxin-transporting cells during embryo development (Steinmann et al., 1999), and PINOID (PID), encoding an auxin-inducible serine–threonine protein kinase, regulates the polar localization of PIN1 in a binary mode of action (Benjamins et al., 2001; Christensen et al., 2000; Friml et al., 2004). The pin1 mutants are defective in PAT and mimic the phenotypic defects induced by treatment with PAT inhibitors (Okada et al., 1991). The pin1 inflorescence terminates in a pin-like structure and the gynoecia of the few flowers formed are trumpet-shaped and lack ovaries (Okada et al., 1991). Mutations in PID also result in severe defects in inflorescence and flower development similar to those seen in pin1 mutants, including distorted apical–basal patterning of the gynoecium (Bennett et al., 1995).
Mutations in the ARF-encoding gene ETTIN (ETT), involved in mediating auxin signals, result in reductions in ovary size, elongation of the gynophore, and development of adaxial tissues in abaxial positions (Sessions and Zambryski, 1995; Sessions et al., 1997). The adaxial–abaxial defects of ett are completely restored by mutations in the bHLH-encoding gene SPATULA (SPT), whereas the apical–basal patterning defects of ett are only partially restored. The adaxial–abaxial defects of ett are due to ectopic expression of SPT, revealing SPT as a downstream target of ETT (Heisler et al., 2001).
We have previously described the isolation and characterization of two Arabidopsis genes, STYLISH1 (STY1) and STYLISH2 ( STY2), which have partially redundant functions in style and stigma development (Kuusk et al., 2002). STY1 and STY2 encode proteins with a putative zinc-binding RING-finger-like domain and belong to a small gene family called the SHI family, with 10 members in Arabidopsis (Fridborg et al., 2001; Kuusk et al., 2006). Apart from the stylar and stigmatic defects, sty1-1 sty2-1 double mutants also display a reduced production of stylar xylem and a basalized point of medial vein bifurcation in the gynoecium (Kuusk et al., 2002). These phenotypes suggest that PAT, auxin metabolism or signalling might be affected by mutations in STY1 and STY2. Here we have investigated the effects of disrupted PAT and auxin signalling in the sty1-1 and sty1-1 sty2-1 mutants. We show that sty1-1, and to a greater degree, sty1-1 sty2-1, confers hypersensitivity to chemical inhibition of PAT in the gynoecium, and enhances the phenotypes of the auxin transport/signalling mutants pin1-5, pid-8 and ett-1. Furthermore, we show that STY1 induces the transcription of an auxin biosynthesis gene, and that auxin homeostasis is altered in plants with altered expression of STY1 and related genes. Together our data suggest that STY1 and the related SHI genes affect various developmental processes by regulating auxin homeostasis.
sty1-1 and sty1-1sty2-1 gynoecia are hypersensitive to NPA
The vascular defects in sty1-1 and sty1-1 sty2-1 gynoecia are similar to those observed in plants treated with PAT inhibitors (Kuusk et al., 2002; Nemhauser et al., 2000), and suggest that auxin homeostasis or signalling might be disturbed in these plants. We therefore examined the response of sty1-1 and sty1-1 sty2-1 to treatment with the PAT inhibitor 1-N-naphtylphtalamic acid (NPA).
Wild-type, sty1-1 and sty1-1 sty2-1 plants were sprayed once with a 100μm NPA solution and the flowers formed post-treatment were observed continuously. In all three genotypes, about 10 flowers per inflorescence showed a phenotype different from mock-treated plants and these flowers were analysed in detail. The NPA-treated gynoecia showing phenotypic alterations were sorted into one of three categories based on the appearance of the valves: reduced, very reduced and valveless. Mildly affected gynoecia, which developed valves that covered more than half the length of the gynoecium (Figure 1b) but still were shorter compared with valves of mock-treated gynoecia (Figure 1a), were termed ‘reduced’. If the valves occupied less than half the length of the organ they were termed ‘very reduced’ (Figure 1c), and the most severely affected gynoecia, completely lacking valves, were termed ‘valveless’ (Figure 1d).
The majority (54%) of the affected NPA-treated wild-type gynoecia were classified as reduced (Figure 1b,k), whereas only 13% were valveless (Figure 1d,k). In contrast, in sty1-1 and sty1-1 sty2-1 plants, valveless gynoecia were found in the majority of flowers formed after NPA treatment (76% and 70%, respectively, Figure 1k). As the response to NPA did not differ significantly between gynoecia of sty1-1 and sty1-1 sty2-1, only NPA-treated sty1-1 sty2-1 gynoecia are shown (Figure1g–j).
The NPA treatment also affected the stylar depression in sty1-1 and sty1-1 sty2-1 gynoecia, which frequently was less deep than in mock-treated sty1-1 and sty1-1 sty2-1 (Figure 1g,h). In valveless sty1-1 and sty1-1- sty2-1 gynoecia, a restored style with a fully developed stigma, lacking any sign of depression in the most apical region, was often formed (Figure 1h). The style region of these gynoecia was morphologically similar to the style region of the wild-type valveless gynoecia (Figure 1d,h). In some sty1-1 sty2-1 flowers, however, the valveless gynoecia displayed a trumpet-shaped apex capped with stigmatic papillae (Figure 1i). In the central region of the stigma of these gynoecia, ovules frequently arose in association with a dome of cells (Figure 1j). Such structures were never observed in NPA-treated wild-type gynoecia of any category. The formation of vasculature in sty1-1, sty1-1 sty2-1 and wild-type gynoecia was similarly affected by the NPA treatment. The point of medial vein bifurcation in each gynoecium was more basalized than in mock-treated gynoecia, whereas the distribution and amount of stylar xylem were not significantly altered (data not shown). Mock-treated wild-type, sty1-1 and sty1-1 sty2-1 plants never showed any of the phenotypes associated with NPA treatment.
sty1-1 enhances the gynoecial defects of the weak pin1-5 mutant
To further study the effects of altered auxin transport in sty1-1 plants, we crossed sty1-1 to the weak pin1-5 mutant. The valves of pin1-5 gynoecia were slightly reduced and often fused along one of the margins (not shown), whereas the style and stigma appeared normal (Figure 2a).
We found that sty1-1 significantly enhanced the phenotype of pin1-5 gynoecia (Figure 2b). Valveless gynoecia, which were never observed in pin1-5, were present in 10% of sty1-1 pin1-5 flowers (Figure 2c).
The apical depression in the style of sty1-1 was less prominent in the pin1-5 background, and a mound of cells, frequently bearing ovules, was formed in a central position of the stigma (Figure 2b). This phenotype was strikingly similar to that seen in some of the valveless, NPA-treated sty1-1 or sty1-1 sty2-1 gynoecia (Figure 1i,j).
sty1-1 and sty1-1 sty2-1 enhance the weak pid-8 mutation
To further investigate the functions of STY1 and STY2 in relation to genes affecting PAT, we crossed sty1-1 and sty1-1 sty2-1 to the weak pid-8 mutant. Alterations in floral organ number are more moderate in pid-8 compared with stronger alleles, but the gynoecial defects are almost as severe (Bennett et al., 1995). In a typical pid-8 gynoecium, the valves were reduced and the gynophore and style were elongated (Figure 3a). However, the phenotype was very variable, ranging from completely valveless (3%) to essentially normal gynoecia (28%; Figure 3e). The flowers of pid-8 mutants had a variable number of sepals, an increased number of petals and a decreased number of stamens compared with wild-type flowers (Figure 3f; Bennett et al., 1995).
The sty1-1 pid-8 and sty1-1 sty2-1 pid-8 lines revealed a synergistic interaction between STY1 and PID. The apical–basal patterning defect of pid-8 mutants was enhanced in double and triple mutant gynoecia. In sty1-1 pid-8 plants, only about 7% of the flowers produced normal gynoecia, compared with 28% in the pid-8 single mutant (Figure 3e). In sty1-1 sty2-1 pid-8, the majority of the gynoecia were valveless (57%; Figure 3d,e). Furthermore, although the pid-8 style and stigma were normal, the apical fusion and proliferation of stylar and stigmatic tissues were more severely reduced in the double mutant compared with sty1-1 (Figure 3a,b). The defects in sty1-1 sty2-1 pid-8 plants were similar to those seen in sty1-1 pid-8, but the triple mutants exhibited a more severe reduction of stylar tissue production (Figure 3c,d).
sty1-1 sty2-1 pid-8 plants showed severe alterations in the number of floral organs produced compared with pid-8 and sty1-1 pid-8 flowers (Figure 3f and data not shown). The number of sepals was decreased, the number of petals increased and the number of stamens was unchanged compared with pid-8 and sty1-1 pid-8 (Figure 3f and data not shown). The changes in sepal and petal number are reminiscent of those found in plants carrying strong pid alleles (Bennett et al., 1995), although these plants develop a more variable number of stamens compared with the sty1-1 sty2-1 pid-8 mutants.
The apical–basal patterning defects in ett-1 gynoecia are enhanced by sty1-1
To investigate whether STY1 modulates the sensitivity to reduced PAT through ETT, we created sty1-1 ett-1 double mutants. Some of the gynoecial defects of ett-1 mutants, such as reduction in ovary size and increased gynophore elongation, were enhanced by the sty1-1 mutation (Figure 4a–c). A number of double mutant gynoecia were nearly completely valveless, and in those, ovules grew on top of a placental surface (Figure 4d). In addition, the apical ends of sty1-1 ett-1 gynoecia were less fused than those of ett-1 and sty1-1, and ovules frequently protruded at the apex (Figure 4b,c). The amount of stylar and stigmatic tissues was reduced compared with ett-1, and the double mutant was sterile. As in ett-1 gynoecia, ectopic tissue of transmitting tract identity was generally present. These results suggest a synergistic interaction between STY1 and ETT during apical–basal patterning and apical fusion of the gynoecium.
We have shown previously that STY1 interacts genetically with SPT (Kuusk et al., 2002). Gynoecia of the sty1-1 spt-2 double mutant were less fused than either single mutant, and no stigmatic papillae were formed (Figure 4g; Kuusk et al., 2002). spt has been reported to be partially epistatic to ett because spt-2 restores the adaxial–abaxial patterning defects of ett-1 and the valves of the double mutant are longer than in the ett-1 single mutant (Figure 4f; Heisler et al., 2001). As in sty1-1 spt-2 and ett-1 spt-2 gynoecia, no stigmatic papillae or abaxial outgrowths of the transmitting tract, respectively, were present in the sty1-1 ett-1 spt-2 triple mutants. However, the triple mutant gynoecia were almost completely unfused (Figure 4h,i), and the valves occupied at least half of each sty1-1 ett-1 spt-2 gynoecium, which was significantly more than in ett-1 or sty1-1 ett-1 (Figure 4c, compare with Figure 4i). This shows that STY1 interacts genetically with ETT and SPT during carpel fusion. STY1 also interacts with ETT during the apical–basal patterning of the gynoecia, whereas spt-2 is epistatic to sty1-1.
STY1 affects the expression of genes involved in the regulation of auxin homeostasis
The STY1 protein contains a putative nuclear localization signal, suggesting that it is active in the nucleus (Fridborg et al., 2001). In an attempt to find down tream targets of STY1, we studied the expression profile of lines carrying an inducible STY1 construct. We used a fusion protein consisting of the full-length STY1 fused to the rat glucocorticoid receptor (GR) domain driven by the cauliflower mosaic virus 35S promoter (35S: STY1–GR; Kuusk et al., 2006). The GR domain makes the protein cytoplasmic, but it is shuttled to the nucleus upon treatment with the synthetic ligand dexamethasone (DEX; Lloyd et al., 1994; Schena et al., 1991). To test whether the STY1–GR fusion protein could complement the sty1-1 mutation, we introgressed the construct into the sty1-1 sty2-1 double mutant. In the absence of DEX, 35S: STY1–GR sty1-1 sty2-1 plants were indistinguishable from sty1-1 sty2-1, whereas, upon DEX application, the stylar phenotype of sty1-1 sty2-1 was restored. An extended DEX treatment resulted in a style phenotype similar to 35S: STY1 plants (Kuusk et al., 2002; data not shown). This suggests that 35S: STY1–GR can complement the sty1-1 mutation in a DEX-dependent manner.
To identify downstream targets of STY1, we performed a microarray experiment comparing global gene expression in 35S: STY1–GR sty1-1 sty2-1 seedlings with and without DEX treatment using CATMA arrays (Allemeersch et al., 2005). RNA was extracted at three time points after DEX treatment (2, 3 and 6 h) in order to differentiate between early- and late-response genes. In all, 88 genes were differentially expressed (q < 0.05, see Experimental procedures), and the majority of these (66) were upregulated. Only 15 genes were significantly altered after 2 h, and all of these were upregulated. After 3 h, 23 genes were significantly upregulated, whereas nine were downregulated. Six hours post-treatment, 56 genes were significantly upregulated and 14 were downregulated. The difference in the number of affected genes at each time point shows that it is possible to distinguish between early- and late-response genes. We examined the identity of all the differentially expressed genes to identify those with a potential role in auxin signalling, transport or metabolism. We found two genes fulfilling these criteria: THREAD/YUCCA4 (YUC4) and GH3-2/YDK1, which both were upregulated (Table 1). YUC4 is the closest homologue of the flavin monooxygenase YUC, involved in auxin biosynthesis (Marsch-Martinez et al., 2002; Zhao et al., 2001), and YDK1 is an auxin-inducible IAA-amido synthetase shown to catalyse the conjugation of IAA to amino acids in vitro (Staswick et al., 2005; Takase et al., 2004). YUC4 was significantly up regulated after 2 h, whereas YDK1 expression was not affected until 3 h after DEX treatment (Table 1).
Table 1. Microarray data from three independent biological replicates showing differential expression of auxin-related genes in 35S: STY1–GR sty1-1 sty2-1 seedlings at various time points after DEX treatment
To confirm the results from the microarray analysis, we performed real-time PCR to monitor the expression of YUC4 and YDK1 at various time points after DEX treatment of 35S:STY1–GR sty1-1 sty2-1 seedlings. This analysis showed that YUC4 was significantly upregulated 30 min after DEX treatment, and that the expression of YDK1 was not significantly altered until 3 h post-treatment (Figure 5). These results are in agreement with the results from the microarray analysis and confirm that YUC4 and YDK1 act downstream of STY1 action, and that YUC4 is induced earlier than YDK1 in DEX-treated 35S: STY1–GR sty1-1 sty2-1 seedlings.
As YUC4 appears to be involved in auxin biosynthesis (Marsch-Martinez et al., 2002) and YDK1 is an auxin-inducible gene likely to be involved in deactivation of the hormone via conjugation (Takase et al., 2004), the elevated YDK1 expression found in DEX-treated 35S: STY1–GR sty1-1 sty2-1 seedlings could be a consequence of increased auxin levels due to YUC4 activation. In order to elucidate this, we examined the expression of the auxin-inducible genes IAA1, IAA2, GH3-1, GH3-3, GH3-5 and GH3-6/DFL1 (Hagen and Guilfoyle, 2002; Nakazawa et al., 2001) using real-time PCR, 3 or 6 h after DEX treatment. We also included GH3-4 in the analysis as it is the closest homologue of YDK1 and has been shown to adenylate IAA in vitro (Staswick et al., 2002, 2005). We detected a significant increase in GH3-3 and IAA1 expression, and GH3-6 and IAA2 showed a reproducible, although not statistically significant, induction after DEX treatment (Table 2). The expression of GH3-5 was significantly reduced, whereas GH3-1 and GH3-4 expression was too low to give reproducible results (Table 2). The expression of IAA1 was also studied 30 min and 1 h after DEX treatment, and like YDK1, no significant upregulation of the IAA1 gene was detected until 3 h after STY1–GR activation (Table 2; data not shown). This shows that a subset of auxin-inducible genes, belonging to different gene families, is upregulated after DEX treatment of 35S:STY1–GR sty1-1 sty2-1 seedlings, suggesting that the levels of active IAA are elevated in these plants.
Table 2. Real-time PCR data from two independent biological replicates showing differential expression of YUC4 and auxin-inducible genes in 35S: STY1–GR sty1-1 sty2-1 seedlings 3a or 6b h after DEX treatment (DEX) and in sty1-1 sty2-1 flower buds (sty1-1 sty2-1)
To elucidate whether the same genes also act downstream of STY1 during flower development, we analysed their expression in wild-type and sty1-1 sty2-1 flower buds of stages 0–12 (stages according to Smyth et al., 1990) using real-time PCR. A reproducible decrease in the expression of YUC4, GH3-2/YDK1, GH3-3, GH3-4, GH3-6/DFL1 and IAA1 was detected in sty1-1 sty2-1 flowers compared with wild-type, although this was not statistically significant for GH3-6/DFL1 and IAA1 (Table 2). The GH3-5 and IAA2 expression was unaffected, and the expression of GH3-1 was too low to allow reproducible results. Taken together, these data suggest that STY1 acts as a positive regulator of YUC4 in both seedlings and flowers, potentially resulting in elevated levels of active auxin, altered expression of auxin-responsive genes and altered auxin homeostasis.
STY1 and related genes affect auxin homeostasis
To more directly assess the effect of STY1 on auxin homeostasis, we measured the levels of free IAA and IAA conjugates in 35S: STY1–GR sty1-1 sty2-1 seedlings 6 h after DEX treatment. DEX induction resulted in a significant increase in the levels of free IAA compared with mock treatment, whereas the levels of the other metabolites measured were not affected (Figure 6a). This confirms that STY1–GR induces the production of active auxin, subsequently resulting in activation of auxin-inducible genes.
To further elucidate the role of STY1 in auxin homeostasis, we measured the levels of free auxin and auxin metabolites in flower buds of wild-type, sty1-1 sty2-1, 35S: STY1, ydk1-D and an SHI family quintuple mutant line combining mutations in STY1 and STY2 with mutations in SHI, LRP and SRS5 ( sty1-1 sty2-1 shi-3 lrp1 srs5-1; Fridborg et al., 2001; Kuusk et al., 2002, 2006). The shi-3, lrp1 and srs5-1 mutations strongly enhance the stylar defects of sty1-1 sty2-1 gynoecia, as the quintuple mutant shows severe reductions in proliferation of apical tissues in the gynoecium, demonstrating a high level of functional redundancy in the SHI gene family during gynoecium development (Kuusk et al., 2006). The activation-tagged line ydk1-D (Takase et al., 2004) was included in the study because YDK1 expression was indirectly induced by STY1–GR in seedlings and ydk1-D is expected to have increased levels of auxin amido-conjugates.
We measured the levels of free IAA as well as the IAA precursor indole-3-acetonitrile (IAN) and the catabolites IAA-aspartate (IAAsp), IAA-glutamate (IAGlu) and oxidized IAA (OxIAA). The IAA metabolite profile differed from wild-type in all analysed genotypes, suggesting that both STY1 and YDK1 affect auxin homeostasis (Figure 6b). The levels of free IAA were decreased compared with wild-type in all tested genotypes, although the differences were not statistically significant. Apart from the reduction in IAA levels, no difference in any other auxin metabolite measured was found in the sty1-1 sty2-1 line (Figure 6b). However, in the sty1-1 sty2-1 shi-3 lrp1 srs5-1 quintuple mutant, the content of all auxin metabolites measured was reduced. Statistically significant reductions were found both for the precursor, IAN, and one of the catabolites, OxIAA (Figure 6b). These results suggest that, when members of the SHI gene family are compromised, auxin biosynthesis is reduced, resulting in a subsequent reduction in auxin catabolism and reduced expression of the YUC4, IAA and GH3 genes. In contrast to the significant increase in IAA level upon transient induction of STY1–GR, constitutive expression of STY1 resulted in reduced levels of IAA, IAN and IAGlu (Figure 6b), suggesting that auxin homeostasis is severely distorted in the 35S:STY1 line. In ydk1-D flower buds, we found, as expected, a reduction in IAA and an increase in IAAsp levels. Furthermore, the concentration of OxIAA was significantly elevated, whereas those of IAGlu and IAN were significantly reduced (Figure 6b). Taken together, these data suggest that STY1 and related genes regulate auxin homeostasis and that the effect is different from that of YDK1.
STY1 and STY2 have previously been shown to influence the development of the style and stigma of the Arabidopsis gynoecium (Kuusk et al., 2002). In this paper, we have shown that STY1, and to some extent STY2, also affect the apical–basal patterning of the gynoecium, which could be the consequence of the role of STY1 as a regulator of auxin homeostasis via the auxin biosynthesis gene YUC4.
sty1-1 enhances the apical–basal patterning defects caused by altered auxin transport or signalling in the gynoecium
PAT is essential for the apical–basal patterning of the gynoecium, and our results show that mutations in STY1 confer hypersensitivity to reductions in PAT during gynoecium development. Despite only subtle apical–basal patterning defects in the gynoecium of sty1-1 plants, that is a slight basalization of the point of medial vein bifurcation (Kuusk et al., 2002), the effects of distorted PAT, mediated by NPA treatment or loss of PIN1 function, in these plants were quite dramatic. sty1-1 pin1-5 plants formed valveless gynoecia, which were never found in pin1-5 single mutants, and sty1-1 and sty1-1 sty2-1 gynoecia were hypersensitive to NPA treatment. Furthermore, sty1-1, and to a greater extent, sty1-1 sty2-1, enhanced the apical–basal patterning defects caused by a mutation in PID, a gene shown to regulate PIN protein localization thereby affecting PAT. Interestingly, a slight reduction of the valves and concomitant elongation of the gynophore occurs when STY1 and at least two additional members of the SHI family are mutated, suggesting that the absence of a detectable external apical–basal defect in sty1-1 sty2-1 gynoecia could be due to functional redundancy within the SHI gene family (Kuusk et al., 2006).
Reduction in the activity of the auxin response factor ETT leads to defects in apical–basal patterning of the gynoecium similar to those observed in plants with inhibited PAT, and ett gynoecia are hypersensitive to reductions in PAT (Nemhauser et al., 2000). Interestingly, sty1-1ett-1 gynoecia had significantly smaller ovaries than the ett-1 counterparts, and sty1-1 thus mimics the effects of reduced PAT in ett-1 gynoecia. This finding, together with the results from the NPA treatment and pin1-5 and pid-8 crosses, suggest that STY1 might be involved in regulating PAT. However, analysis of PIN1:PIN1–GFP expression in gynoecia of wild-type and sty1-1 sty2-1 did not reveal any clear difference in PIN1–GFP protein level or localization (our unpublished results). Furthermore, the apical–basal patterning defects of ett mutants are also enhanced by mutations in the transcriptional co-factor SEUSS (SEU), which interacts physically with ETT and is involved in auxin signalling (Pfluger and Zambryski, 2004), implying that STY1 could regulate auxin signalling equally well as auxin transport.
We have previously shown that STY1 interacts genetically with SPT, which is known to be directly or indirectly regulated by ETT at the transcriptional level (Heisler et al., 2001; Kuusk et al., 2002). spt-2 is partially epistatic to ett-1 in the apical-basal patterning of the gynoecium, and spt mutants exhibit reduced sensitivity to NPA treatment, indicating that interpretation of the auxin signal in the gynoecium is SPT-dependent (Heisler et al., 2001; Nemhauser et al., 2000). We found that sty1-1 enhanced the apical–basal patterning defect of ett-1 gynoecia, but the effect was masked in sty1-1 ett-1 spt-2 triple mutants. This epistasis of spt-2 over sty1-1 indicates that STY1 and SPT regulate apical–basal patterning in the same pathway.
STY1 and related genes affect auxin homeostasis
Our microarray and real-time PCR analyses clearly show that YUC4 and several auxin-inducible genes act downstream of STY1 gene function. They further indicate that STY1 could act as a transcriptional activator of YUC4 because increased expression of this gene was detected very early (30 min) after DEX treatment of 35S:STY1–GR sty1-1 sty2-1 seedlings. YUC4 is the closest homologue of the auxin biosynthesis gene YUC, and has been shown to confer similar auxin overproduction phenotypes as YUC when overexpressed (Marsch-Martinez et al., 2002). It is therefore likely that YUC4 performs the same biochemical reaction as YUC in auxin biosynthesis. The STY1–GR-induced expression of auxin-inducible genes occurred approximately 3 h later than the induction of YUC4, implying that STY1–GR activates their expression through a YUC4-mediated elevation of auxin levels. Accordingly, the increased expression of auxin-inducible genes correlated with a significant increase in the level of free IAA in DEX-treated 35S: STY1–GR sty1-1 sty2-1 seedlings, suggesting that STY1 is a positive regulator of auxin biosynthesis. Of the auxin-inducible GH3-like genes studied, YDK1 and GH3-3 were activated by STY1–GR after DEX treatment, whereas GH3-5 and -6 were not. This difference in GH3 gene responsiveness to STY1–GR is not unexpected because available data reveal that GH3 genes differ in their sensitivity to auxin and/or in their spatial expression. For example, YDK1 and GH3-3 but not GH3-5 or -6 are upregulated in rosette leaves of plants expressing a microRNA-resistant version of ARF17 (Mallory et al., 2005). Furthermore, auxin-induced expression of YDK1, GH3-1, -3 and -4 is four times more elevated compared with GH3-5 and -6 according to microarray data (GEO accession GSM9969, http://www.ncbi.nlm.nih.gov/geo/). This suggests that induction of STY1–GR activity results in auxin levels sufficient for the induction of YDK1 and GH3-3 but not of GH3-5 and -6.
Using real-time PCR, we could show that the expression of YUC4 and auxin-inducible genes was also affected in sty1-1 sty2-1 flower buds, suggesting that STY1 regulates auxin homeostasis not only in seedlings, or in conditions when STY1 is overexpressed, but also during flower development. However, when measured directly, the level of free IAA was only slightly reduced in sty1-1 sty2-1 flower buds, and the level of precursor and catabolites was unaffected. The moderate effect of loss of STY1 and STY2 on IAA levels is not surprising considering the high degree of functional redundancy among the SHI genes and the limited expression domains of these genes (Fridborg et al., 2001; Kuusk et al., 2002, 2006). In flower buds of the sty1-1 sty2-1 shi-3 lrp1 srs5-1 quintuple mutant, the levels of most IAA metabolites measured, including free IAA, were reduced, suggesting that the SHI family genes regulate auxin homeostasis in a redundant manner.
Several mechanisms that act to maintain auxin homeostasis have been described. When auxin catabolism is increased, biosynthesis is expected to be activated to counteract the reduced auxin levels. If, on the other hand, auxin biosynthesis is reduced, a reduced catabolism of free IAA, moderating the effects of the reduced production, would be expected. The reduced levels of almost all measured auxin metabolites in sty1-1 sty2-1 shi-3 lrp1 srs5-1 flower buds, together with the reduced expression of YUC4 in sty1-1 sty2-1, are therefore in accordance with a role of STY1 and related genes in regulating auxin biosynthesis. Constitutive expression of STY1, however, appears to activate several pathways modulating both auxin synthesis and catabolism, and most likely does not accurately reflect the role of STY1 in the wild-type.
In the ydk1-D mutant, we expected to find an increase in the levels of IAA conjugates and potentially an increase in some auxin precursors. In line with these expectations, we found an increase in the catabolites IAAsp and OxIAA but a decrease in IAGlu. This suggests that auxin catabolism is increased in these plants and indicates that YDK1 might have some specificity with regard to amino acid substrate. Furthermore, the level of the precursor IAN was significantly reduced, which is somewhat surprising. It might, however, be due to an increased flux from IAN to IAA, to maintain homeostasis, without a concomitant increase in the synthesis of IAN.
Altered auxin homeostasis could cause the sty1-1 sty2-1 phenotype
According to the hypothesis proposed by Nemhauser et al. (2000), auxin is synthesized apically and transported basipetally through the gynoecium. High auxin levels would specify style/stigma, intermediate concentrations would give rise to the ovary, and the gynophore would result from low or no auxin. If auxin promotes the development of the style and stigma, as stated by the model, our results suggest that STY1 and STY2 could contribute to the formation of style and stigma through activation of auxin biosynthesis, and that the stylar defects observed in sty1-1 and sty1-1 sty2-1 plants are due to reduced auxin levels. If so, increased auxin levels in the apical parts of sty1-1 and sty1-1 sty2-1 gynoecia would be predicted to restore the stylar phenotype of these plants. We found indirect evidence to suggest that this is the case. It has been proposed that treatment with NPA causes accumulation of auxin at the sites of synthesis, hence PAT inhibition in the gynoecium most likely leads to an increase in auxin levels in the apical parts (Nemhauser et al., 2000). In accordance with this hypothesis, the stylar defects of sty1-1 and sty1-1 sty2-1 plants were restored by NPA, which implies that the reduced style proliferation in these plants is due to reduced auxin levels.
STY1 and STY2 expression in the gynoecium is confined to the most apical part, and STY1 transcript is already present from stage 6 of flower development, whereas activity of the STY2 promoter is first detected during stage 9 (Kuusk et al., 2002). The temporally and spatially restricted expression of STY1 and STY2 is in agreement with their role in style and stigma morphogenesis, but suggests that their effect is not cell-autonomous in the apical–basal patterning of the gynoecium. If the hypothesis proposed by Nemhauser et al. (2000) is correct, reduced auxin biosynthesis in the apical part of sty1-1 and sty1-1 sty2-1 gynoecia would affect not only style morphogenesis, but also the amount of auxin transported basipetally, and could explain the defects in apical–basal patterning in these plants. The apical–basal defects observed in sty1-1 and sty1-1 sty2-1 plants are, however, very subtle, which could be the consequence of functional redundancy between members in the SHI gene family, as mentioned previously, and the capacity of the auxin transport system to buffer changes in auxin levels. The latter has been shown to occur in the embryo, in which alterations in auxin concentration cause no apparent developmental defects but render the embryos more sensitive to PAT inhibition (Weijers et al., 2005). It is not unlikely that a similar PAT-dependent buffering mechanism of changes in auxin metabolism is also present in the gynoecium, and this could explain the increased sensitivity of sty1-1 and sty1-1 sty2-1 gynoecia to alterations in PAT.
Taken together, our results indicate that YUC4 is a primary target of STY1 action, and that STY1 thus acts as an activator of auxin biosynthesis. This hypothesis is supported by the altered expression of auxin-inducible genes and changes in auxin homeostasis detected in DEX-treated 35S: STY1 sty1-1 sty2-1 seedlings and sty1-1 sty2-1 flower buds. Furthermore, the hypersensitivity of sty1-1 sty2-1 gynoecia to altered PAT and auxin signalling and the rescue of the sty1-1 sty2-1 stylar defects through NPA treatment suggest that reduced auxin biosynthesis could cause the developmental aberrations in sty1-1 and sty1-1 sty2-1 gynoecia. We therefore propose that one primary role of STY1, and additional SHI family genes, is regulation of auxin biosynthesis, and that this causes at least some of the developmental defects observed when these genes are mutated.
Plant material and genetics
Arabidopsis wild-type accession Columbia (Col) was used unless otherwise stated. sty1-1, sty2-1 and sty1-1 sty2-1 mutants and the 35S: STY1 line are in the Col background and have been described previously (Kuusk et al., 2002). The sty1-1 sty2-1 shi-3 lrp1 srs5-1 quintuple mutant is described by Kuusk et al. (2006). ett-1 and pid-8 mutant seeds were provided by the Arabidopsis Biological Research Centre (ABRC, Columbus, OH, USA). spt-2, pin1-5 and ydk1-D seeds were kindly provided by John Bowman, School of Biological Sciences, Monash University, Melbourne, VIC 3800, Australia, David Smyth, School of Biological Sciences, Monash University, Australia and Minami Matsui, Genomic Sciences Center, RIKEN, Japan, respectively. pid-8 and ett-1 are in Wassilevskija, whereas spt-2 and pin1-5 are in Landsberg erecta. The pid-8 mutant was crossed with Col, and the F3 generation of this cross was used in the experiments. Double mutant lines were generated by cross-fertilization of homozygous single mutant lines. Double mutants were identified in the F2 population by their phenotype, and for sty1-1 and sty2-1 with PCR. The genotypes were confirmed by segregation analysis in the F3 generation. The sty1-1 pid-8 double mutant was backcrossed twice to sty1-1, and F3 double mutant offspring were used in all studies. For DEX treatment, 35S: STY1–GR sty1-1 sty2-1 plants were grown in liquid culture (0.5x MS medium with 2% sucrose) with gentle agitation.
Plants were sprayed with a solution containing 0.01% Silwet L-77 (Lehle Seeds, Round Rock, TX, USA) and 100 μm 1-N-naphtylphtalamic acid (NPA; Riedel-de-Haen, Seelze, Germany) or a control solution with 0.01% Silwet L-77 according to the method described by Nemhauser et al. (2000). Flowers were checked regularly, and affected flowers were seen 12–18 days post-treatment. The experiment was repeated twice with similar results. Results shown are from one experiment.
RNA was isolated from 14-day-old 35S:STY1–GR sty1-1 sty2-1 seedlings treated with 10 μm dexamethasone (Sigma-Aldrich, Schnelldorf, Germany) or with a mock solution (0.1% EtOH), and from sty1-1 sty2-1 or Col inflorescences including flowers of stages 0–12. Total RNA was extracted using the RNeasy Plant Mini Kit (Qiagen, Valencia, CA, USA) according to the manufacturer's instructions. Aliquots (5 μg) of total RNA were reverse- transcribed to cDNA using an oligo(dT) 20-mer with a T7 promoter in the 5′ position and two randomized nucleotides in the 3′ position, using Superscript III reverse transcriptase (Invitrogen, Carlsbad, CA, USA).
The CATMA microarray used in the presents study comprised 21 120 gene-specific sequence tags (GSTs) for annotated Arabidopsisthaliana genes (Hilson et al., 2004).
Probe labelling and microarray hybridization protocols
Data acquisition, normalization and statistical analysis
Hybridized arrays were scanned with an Axon 4000B scanner using the GenePix 3.1 analysis software (Axon Instruments, Foster City, CA, USA). The GenePix 3.1 software was also used to quantify spot intensities as well as the local background intensities. All subsequent data analysis was performed in R (http://www.r-project.org) using marray and limma in the open source Bioconductor project (http://bioconductor.org) and Cyber-T (Baldi and Long, 2001). Data points were removed if a spot was flagged during data acquisition (e.g. as a bad spot, absent spot, etc.) or if the spot intensity was below a threshold determined by a model-based statistical method, based on the standard deviation of background differences between a spot and the neighbouring spots (Yang et al., 2001). Spot intensities were then background-corrected using the normexp option (offset = 50) in limma. The data were normalized using within-print-tip-group intensity-dependent normalization with the marray package. To identify differentially expressed genes, we used a regularized t-test that combines information from gene-specific and global average variance estimates by using a weighted average of the two as the denominator for a gene-specific t-test (Baldi and Long, 2001). The obtained P-values were corrected for multiple testing based on the positive false discovery rate (pFDR; Storey, 2002) by calculating the q value, which is the pFDR analogue of the P-value (Storey, 2002). Microarray analysis was performed on three independent biological samples.
Quantitative real-time PCR
The cDNA was diluted to 2 ng/μl, and 5 μl of the diluted cDNA was used as a template for quantitative real-time PCR analysis. The cDNA was amplified using the SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA, USA) on an ABI 7000 thermocycler (Applied Biosystems). Primers for the ACTIN2 gene were used as an internal control to normalize the expression data for each gene. For primer sequences used in this study, see Table S1. The PCR conditions were as follows: 50°C for 2 min and 95°C for 10 min, followed by 40 cycles of 95°C for 15 sec, 60°C for 1 min. A dissociation kinetics analysis was performed at the end of the experiment to check the specificity of annealing. Three replicates were performed for each sample in each experiment. The experiments were repeated twice independently. Results were analysed according to the method described by Muller et al. (2002).
Auxin metabolism analysis
Inflorescences containing flower buds of stages 0–12 were collected and frozen in liquid nitrogen. Samples were collected from three independent biological replicates.
Plant material (20–100 mg) was mixed with 1 ml of 50 mm sodium phosphate buffer, pH 7, containing a mixture of [indole-13 C6] internal standards (2 ng of OxIAA and IAA, 600 pg of IAAsp and IAGlu, 100 pg IAAla and IALeu, 50 ng IAN per sample), homogenized in a Retsch Mixer Mill (Retsch GmbH, Haan, Germany), and extracted for 1 h at 4°C. Samples were then centrifuged (20800 g for 15 min at 4°C), and supernatants transferred to clean tubes and acidified to pH 2.7 with 1 m hydrochloric acid. Solid-phase extraction was performed using 30 mg Waters Oasis MAX columns (Waters Corp., Milford, MA, USA). After sample application, columns were washed with 1 ml of 10 mm ammonium hydroxide. The neutral fraction (containing IAN) was removed from the columns with 1 ml acetone, and then the acidic fraction (containing IAA, OxIAA and the amide conjugates) was eluted using 1 ml of acetonitrile containing 2% v/v formic acid.
Acidic fractions were vacuum-dried, dissolved in 25 μl methanol, and reacted with 225 μl diazomethane in diethylether for 20 min. The excess of derivatization reagent was quenched with 100 μl of 2 m acetic acid in n-heptane, and samples were dried in a stream of nitrogen. Samples from both neutral and acidic fractions were reconstituted with 10 μl of 20% methanol prior to analysis by LC-MS/MS.
For liquid chromatography, a 10 × 1 mm Thermo BetaMax precolumn (Thermo Electron Corp, Waltham, MA, USA) connected to a 50 × 1 mm Waters Symmetry Shield C-18 analytical column was used. A linear gradient of 20–90% methanol containing 1% v/v formic acid over 25 min at a flow rate of 35 μl/min was employed to separate analytes of interest, followed by 5 min of washing with 100% methanol and a 15 min 20% methanol/1% formic acid equilibration period for each injection. The Waters Quattro Ultima mass spectrometer was operated in MRM mode in conditions essentially as described previously (Kowalczyk and Sandberg, 2001), except that source block and desolvation temperatures were decreased to 100 and 290°C, respectively. For IAN quantification, the collision energy was decreased to 16 eV. Acquired data were processed using Waters MassLynx software.
We thank Gun-Britt Berglund, Cecilia Wärdig and Staffan Tjus for skilful technical assistance, Stefan Gunnarsson and Gary Wife for providing excellent SEM facilities, and Pelle Larshammar for setting up the real-time PCR. We are grateful to Ingela Fridborg, Folke Sitbon and Jens Sundström for helpful comments on the manuscript. We acknowledge the Wallenberg Consortium North for providing the CATMA chips and for assistance in the production of transgenic plants via the Arabidopsis model organism platform in Uppsala. This work was supported by grants from the Swedish Research Council, the Swedish Research Council for Environment, Agricultural Sciences and Spatial Planning, and the Nilsson-Ehle foundation.