Auxin and Ethylene Induce Flavonol Accumulation through Distinct Transcriptional Networks (original) (raw)

• Journal List
• Plant Physiol
• v.156(1); 2011 May
• PMC3091047

Plant Physiol. 2011 May; 156(1): 144–164.

Department of Biology, Wake Forest University, Winston-Salem, North Carolina 27109 (D.R.L., G.K.M.); Department of Biological Sciences (M.V.R., P.V., B.S.J.W.) and Department of Biochemistry (W.K.R., R.F.H.), Virginia Tech, Blacksburg, Virginia 24061; and Department of Botany, University of Wisconsin, Madison, Wisconsin 53706 (N.D.M.)

1This work was supported by the National Science Foundation Arabidopsis 2010 Program (grant nos. IOB–0820717 to G.K.M. and 0820674 to B.S.J.W. and R.F.H.) and National Science Foundation Molecular Biochemistry (grant no. MCB–0445878 to B.S.J.W.), by the U.S. Department of Agriculture-National Research Initiative Competitive Grants Program (grant no. 2006–03406 to G.K.M.), by the National Science Foundation Major Research Instrumentation Program for purchase of the confocal microscope (grant no. MRI–0722926 to Anita McCauley and G.K.M.), and by the National Science Foundation Plant Genome Program (grant no. DBI–0621702 to Edgar Spalding).

2Present address: Mycobacteria Research Laboratories, Department of Microbiology, Immunology, and Pathology, Colorado State University, Fort Collins, CO 80523.

The authors responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) are: Brenda S.J. Winkel (ude.tv@lekniw) and Gloria K. Muday (ude.ufw@yadum).

[C]Some figures in this article are displayed in color online but in black and white in the print edition.

[W]The online version of this article contains Web-only data.

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Received 2011 Jan 18; Accepted 2011 Mar 19.

Copyright © 2011 American Society of Plant Biologists

Abstract

Auxin and ethylene are key regulators of plant growth and development, and thus the transcriptional networks that mediate responses to these hormones have been the subject of intense research. This study dissected the hormonal cross talk regulating the synthesis of flavonols and examined their impact on root growth and development. We analyzed the effects of auxin and an ethylene precursor on roots of wild-type and hormone-insensitive Arabidopsis (Arabidopsis thaliana) mutants at the transcript, protein, and metabolite levels at high spatial and temporal resolution. Indole-3-acetic acid (IAA) and 1-aminocyclopropane-1-carboxylic acid (ACC) differentially increased flavonol pathway transcripts and flavonol accumulation, altering the relative abundance of quercetin and kaempferol. The IAA, but not ACC, response is lost in the transport inhibitor response1 (tir1) auxin receptor mutant, while ACC responses, but not IAA responses, are lost in ethylene insensitive2 (ein2) and ethylene resistant1 (etr1) ethylene signaling mutants. A kinetic analysis identified increases in transcripts encoding the transcriptional regulators MYB12, Transparent Testa Glabra1, and Production of Anthocyanin Pigment after hormone treatments, which preceded increases in transcripts encoding flavonoid biosynthetic enzymes. In addition, myb12 mutants were insensitive to the effects of auxin and ethylene on flavonol metabolism. The equivalent phenotypes for transparent testa4 (tt4), which makes no flavonols, and tt7, which makes kaempferol but not quercetin, showed that quercetin derivatives are the inhibitors of basipetal root auxin transport, gravitropism, and elongation growth. Collectively, these experiments demonstrate that auxin and ethylene regulate flavonol biosynthesis through distinct signaling networks involving TIR1 and EIN2/ETR1, respectively, both of which converge on MYB12. This study also provides new evidence that quercetin is the flavonol that modulates basipetal auxin transport.

Auxin and ethylene have long been known to be key regulators of plant growth and development. Recent genetic evidence suggests that a variety of root developmental processes, including elongation, gravitropism, and lateral root formation, are regulated by cross talk between these two hormones (Rahman et al., 2001; Buer et al., 2006; Růzicka et al., 2007; Stepanova et al., 2007; Swarup et al., 2007; Negi et al., 2008, 2010). The ethylene- and auxin-signaling pathways have been extensively studied, and studies employing global gene expression analyses and the unique genetic tools available in Arabidopsis (Arabidopsis thaliana) have identified a number of the key signaling proteins as well as the transcriptional response targets (Kendrick and Chang, 2008; Chapman and Estelle, 2009).

For ethylene signaling, the family of receptors that initiate the ethylene response cascade includes the well-characterized ethylene receptor, Ethylene Resistant1 (ETR1; Chang et al., 1993). Constitutive Triple Response1 (CTR1), an active protein kinase with sequence similarity to the catalytic domain of RAF protein kinase, is downstream from ETR1 and believed to function as a negative regulator of signaling (Kieber et al., 1993; Huang et al., 2003). Ethylene Insensitive2 (EIN2) is a membrane protein whose specific biochemical activity is not yet clear, although it acts downstream from both ETR1 and CTR1. Null mutations in EIN2 result in complete loss of ethylene responsiveness, suggesting that it is an essential, positive modulator of ethylene signaling (Alonso et al., 1999). Further downstream from these signaling proteins are a range of transcription factors, including EIN3 and Ethylene Response Factor1 (ERF1), which control the synthesis of proteins that mediate physiological and developmental responses to ethylene (Kendrick and Chang, 2008).

In auxin signaling, Transport Inhibitor Response1 (TIR1) encodes an auxin receptor, an E3-ubiquitin ligase, which functions to degrade transcriptional repressors and thereby induce gene expression (Dharmasiri et al., 2005; Kepinski and Leyser, 2005). These negative transcription factors are encoded by the AUXIN/INDOLE-3-ACETIC ACID (AUX/IAA) gene families (Ulmasov et al., 1987), and upon their degradation, Auxin Response Factor (ARF) family proteins are released from complexes and generally act as positive transcription factors enhancing the expression of target genes, which then affect growth and developmental responses (Ulmasov et al., 1999; Shin et al., 2007).

Yet, the cross talk between the auxin- and ethylene-responsive transcriptional networks has received limited study (Stepanova and Alonso, 2009). One recent genome-wide transcriptional analysis of dark-grown seedlings found only 191 genes whose expression was affected by both auxin and ethylene, which represented 27% and 18% of all the ethylene- and auxin-regulated genes, respectively (Stepanova et al., 2007). Using ein2-5 and auxin resistant1 (aux1) mutants to selectively block ethylene and auxin responses, groups of genes were identified that reciprocally regulate each other’s biosynthesis, influence each other’s response pathways, or act independently on the same target genes (Stepanova et al., 2007). Yet, the functional significance of the transcripts falling into these three classes is not well understood, and there is little information on the ultimate target genes that then control plant growth and development. Genes encoding enzymes of flavonoid metabolism may be among these ultimate targets of auxin and ethylene transcriptional networks. Previous studies have suggested that flavonol accumulation may be induced by both auxin and ethylene and have implicated these specialized metabolites in hormone-dependent developmental pathways (Buer and Muday, 2004; Buer et al., 2006) but have not examined whether this accumulation is transcriptionally regulated.

Flavonols have been shown to regulate auxin transport and dependent physiological processes, including root gravitropism and branching (Brown et al., 2001; Buer and Muday, 2004; Peer et al., 2004; Buer and Djordjevic, 2009). Quercetin and, to a lesser extent, kaempferol inhibit auxin efflux from preloaded hypocotyl segments, suggesting that specific flavonols function as auxin transport inhibitors in hypocotyl tissue (Jacobs and Rubery, 1988). Auxin transport is elevated in inflorescences, hypocotyls, and roots of plants with the transparent testa4-2 (tt4-2) mutation, which make no flavonoids (Murphy et al., 2000; Brown et al., 2001). A comparison of the root gravitropic responses of the wild type and several tt4 alleles identified a delay in root gravitropism when flavonoid synthesis is abolished, which is reversed by chemical complementation by naringenin (Buer and Muday, 2004; Buer et al., 2006). Flavonoids promote gravitropism presumably by regulating auxin movement in the root tip, which modulates differential growth (Buer and Muday, 2004). Finally, factors that regulate flavonoid biosynthesis, such as light levels (Jensen et al., 1998; Rashotte et al., 2003), wounding and pathogen attacks (Mathesius et al., 1998; Berleth and Sachs, 2001), and gravity stimulation (Buer and Muday, 2004; Buer et al., 2006), also affect auxin transport. Flavonoid synthesis is regulated by a variety of developmental and environmental cues (Winkel-Shirley, 2002). The expression of genes in this pathway is light dependent (Pelletier and Shirley, 1996; Cain et al., 1997) and regulated by light quality and quantity through distinct photoreceptors (Hemm et al., 2004). Among the transcription factors tied to light signaling that control this process are Elongated Hypocotyl, which has a unique role in cross talk between auxin and light signaling in both roots and shoots of Arabidopsis (Cluis et al., 2004; Sibout et al., 2006). Taken together, these results suggest that flavonoids may be important regulators of plant growth and development in response to auxin and ethylene signaling and that these compounds may act by modulating auxin transport.

The general phenylpropanoid pathway produces important specialized metabolites, including structural components such as lignin (Boerjan et al., 2003; Vogt, 2010), flavonoids that prevent photodamage (Li et al., 1993; Bharti and Khurana, 1997; Bieza and Lois, 2001), and flavonols that regulate plant growth and development (Brown et al., 2001; Buer and Muday, 2004; Peer et al., 2004). A representation, shown in Figure 1, depicts the order of enzymes and intermediates that make up the core of the flavonoid biosynthetic pathway (Winkel-Shirley, 2001; Koes et al., 2005). Chalcone synthase (CHS) catalyzes the first committed step in this pathway, and Arabidopsis tt4-2 null mutants, which have lesions at this locus, accumulate no downstream compounds (Shirley et al., 1995; Burbulis et al., 1996; Saslowsky et al., 2000). Another important step is controlled by the branch point enzyme, flavonoid 3′-hydroxylase (F3′H), which converts dihydroxykaempferol to dihydroxyquercetin and is encoded by the TT7 locus in Arabidopsis (Schoenbohm et al., 2000). In addition to the synthesis of the core aglycone flavonols outlined in this figure, flavonols can be extensively glycosylated to regulate both their structure and function (Winkel-Shirley, 2001). In young Arabidopsis seedlings, aglycone flavonols are of low abundance or nondetectable, as determined by HPLC analyses (Burbulis et al., 1996; Böttcher et al., 2008; Yonekura-Sakakibara et al., 2008).

The flavonoid biosynthetic pathway. Intermediate compounds and enzymes in the pathway are shown with mutant alleles represented in brackets. The structures of naringenin and quercetin are shown, and downstream products are represented at the bottom of the diagram. Enzymes, mutants, and metabolites central to this study are shown in boldface. This figure is modified from Buer and Muday (2004).

A number of transcription factors that control the expression of the genes encoding flavonoid biosynthetic enzymes have been identified (Stracke et al., 2010b). The current model of the transcriptional regulation of the flavonoid biosynthetic pathway is that different R2R3-MYB, basic helix-loop-helix, and WD40-type transcription factors control expression of the pathway in various tissues in response to a broad set of stimuli (Stracke et al., 2007; Gonzalez et al., 2008; Feyissa et al., 2009). These transcriptional regulators include Production of Anthocyanin Pigment (PAP1/MYB75) and Transparent Testa Glabra1 (TTG1; a WD40 protein), which control the accumulation of proanthocyanidin and anthocyanin, by regulation of targets including Dihydroflavonol 4-Reductase (DFR) and BANYULS (Shirley et al., 1995; Baudry et al., 2004). In contrast, flavonol biosynthetic genes have recently been shown to be controlled in tissue- and cell type-specific patterns by three MYB proteins that appear to function without a basic helix-loop-helix partner (Stracke et al., 2007). Of these, MYB12 specifically activates flavonol genes in Arabidopsis roots with only minor contributions from MYB11 and MYB111 (Stracke et al., 2007).

This study quantified the hormonal regulation of the flavonoid biosynthetic pathway by analyzing changes in gene expression, enzyme accumulation, and accumulation of flavonols with high spatial and temporal resolution. We employed mutant analyses to determine the necessary molecular signaling and regulatory components and found evidence for distinct TIR1- and ETR1-mediated signaling pathways controlling responses to auxin and ethylene with a consensus role of MYB12 in both signaling networks. Collectively, these experiments provide insight into the mechanisms by which auxin and ethylene induce the expression of genes required for the synthesis of these important specialized metabolites, forging new connections between the auxin and ethylene signaling pathways and flavonoid regulatory and structural target genes. This work further provides new support for the primary role of quercetin and its derivatives in modulating root basipetal (or shoot-ward) auxin transport and dependent physiological processes. Thus, one of the outcomes of auxin signaling is a transcriptional activation of flavonol biosynthesis, leading in turn to modulation of auxin transport.

RESULTS

Auxin and Ethylene Induce Genes Encoding Flavonoid Biosynthetic Enzymes

To examine the effects of auxin and ethylene on flavonoid metabolism, the accumulation of mRNA encoding enzymes in the flavonoid pathway in seedling primary roots was examined by quantitative real-time (qRT)-PCR analysis of reverse-transcribed total RNA. The RNA was extracted from roots of seedlings grown on clear agar substrate, exposing both roots and shoots to light. To study root development and its hormonal controls, most investigators grow seedlings on nutrient medium in petri dishes, such that both roots and shoots are exposed to light (Brady et al., 2007; Péret et al., 2009). Under these conditions, there is robust expression of genes of flavonoid metabolism (Brady et al., 2007) and well-characterized hormonally regulated gene expression and developmental mechanisms, including auxin-ethylene cross talk (Růzicka et al., 2007; Stepanova et al., 2007; Swarup et al., 2007). The abundance of transcripts for CHS, FLAVONOL SYNTHASE (FLS), and F3′H was analyzed in cDNA prepared from roots of untreated and hormone-treated seedlings. The levels of these transcripts relative to those of actin (ACT2) were normalized to an untreated control after a 6-h treatment with IAA or 1-aminocyclopropane-1-carboxylic acid (ACC), as shown in Figure 2. Both 1 μm IAA and 1 μm ACC led to a greater than 3-fold induction of CHS and FLS mRNA accumulation, which was statistically significant (P < 0.04). _F3′H_ was significantly induced by IAA (_P_ < 0.01) but not by ACC (_P_ > 0.9). We also examined the transcript abundance of genes encoding additional enzymes in the flavonoid biosynthetic pathway, including CHALCONE ISOMERASE (CHI) and FLAVANONE 3 HYDROXYLASE (F3H), and DFR with and without IAA and ACC treatment. We treated wild-type seedlings with both IAA and ACC and quantified the abundance of CHS, F3′H, and FLS transcripts, as shown in Figure 2. For CHS and FLS, the effects of both hormones together was additive, while for F3′H, the transcript abundance was unchanged by the addition of ACC in either the absence and presence of IAA. In Table I, the abundance of each transcript relative to the internal actin standard is reported as well as the fold induction 4 h after IAA or ACC treatment. CHI showed a similar induction to CHS, while little or no change was observed for F3H in response to either treatment. Although it is surprising that F3H transcripts were unchanged by these treatments, the abundance of these transcripts was substantially higher than those encoding many of the other pathway enzymes, suggesting that F3H protein levels might not limit pathway function. The low level of F3′H message relative to actin and its position in the metabolic pathway is consistent with the enzyme encoded by this transcript acting as an important metabolic branch point. DFR message was not detected in the cDNA samples used for this study, using four independent primer sets, all of which were able to detect DFR in genomic DNA samples (data not shown). The absence of DFR message in Arabidopsis roots, even upon exposure to light, is consistent with published microarray data and data deposited in the Gene Expression Omnibus database (Barrett et al., 2009) and with the lack of detectable anthocyanin accumulation in this tissue. Overall, the transcript abundance values of untreated control roots correlate well with publicly available expression data (Brady et al., 2007) and suggest cases where lower transcript abundance of genes encoding certain pathway enzymes may lead to low levels of enzymes that limit the activity of biosynthetic pathway steps.

IAA and ACC enhance the expression of genes encoding flavonol pathway enzymes. The abundance of CHS, FLS, and F3′H transcripts relative to actin and normalized to the untreated wild type (WT) 6 h after treatment with 1 μm IAA and ACC is shown in the wild type, tir1, ein2, etr1, and myb12. The basal level and fold induction of CHS (A), FLS (B), and F3′H (C) transcript abundance after treatment with IAA and ACC are shown in all genotypes. Mean and se of three biological replicates are shown. * Significant difference (P < 0.05) between the mutant and the wild type within a treatment; # significant difference between treated and untreated controls within a genotype as determined by Student’s t test (P < 0.05).

Table I.

Ratios of target gene transcript to actin in control, IAA-, and ACC-treated wild-type samples as determined by qRT-PCR

Roots were transferred to control medium or medium containing 1 μm ACC or IAA for 4 h. RNA was harvested from roots and amplified by qRT-PCR with gene-specific primers. The average ± se of three biological replicates is reported. Fold change, shown in parentheses, was determined by dividing the target actin ratios of the treated sample by the control ratio. ND, Not detected.

We examined the genetic controls of IAA and ACC that induced flavonol synthesis by examination of the transcript abundance of these pathway genes in tir1-1 or ein2-5 and etr1-3, which are mutants defective in auxin or ethylene signaling, respectively (Stepanova and Ecker, 2000; Dharmasiri et al., 2005), in the presence and absence of both IAA and ACC. Untreated tir1-1 seedlings exhibited significantly lower CHS and F3′H transcript accumulation in roots than did the wild type (P < 0.04), and _CHS_ and _FLS_ were unchanged by IAA treatment (_P_ > 0.09). However, IAA treatment caused a significant 2.0-fold increase in F3′H transcript abundance in tir1-1 (compared with a 3.8-fold increase in the wild type; P < 0.02). This induction brought _F3′H_ transcript abundance in _tir1-1_ to a level comparable to that of the untreated wild type (_P_ > 0.11). As expected, the response of CHS and FLS to ACC was not significantly affected by the tir1-1 mutation, consistent with the existence of independent ethylene and auxin signaling pathways.

CHS and FLS transcripts were reduced by 39% and 35%, respectively, in untreated ein2-5 relative to untreated control (P < 0.05). In _etr1-3_, the magnitude of ACC-enhanced accumulation of _CHS_ and _FLS_ transcripts was lower than in the wild type (Fig. 2), but transcript abundance of these genes in untreated _etr1-3_ was not significantly different from the wild type (_P_ < 0.08). This is consistent with previous reports that _etr1-3_ retains some ethylene signaling activity (Hall et al., 1999). There were no ACC-induced changes in _CHS_, _FLS_, or _F3′H_ transcript abundance in _ein2-5_ or _etr1-3_ relative to untreated controls (_P_ > 0.2). In contrast, auxin induction of CHS, FLS, and F3′H was intact in ein2-5 and etr1-3. Together with the additive effect of cotreatment with IAA and ACC on the abundance of flavonoid enzyme mRNA, these results indicate that well-defined auxin and ethylene signaling pathways regulate the synthesis of flavonoid metabolic enzymes and that the signaling pathways for IAA- and ACC-induced flavonol accumulation are distinct and nonoverlapping.

The effect of mutations in MYB12 on baseline expression of these three genes as well as their induction by IAA and ACC was also examined. Untreated myb12 exhibited a nonsignificant (P > 0.1) reduction in CHS, F3′H, and FLS mRNA abundance. However, when myb12 was treated with either IAA or ACC, all three of these transcripts did not significantly change (P > 0.5). These data are consistent with a requirement for MYB12 for the hormonally enhanced expression of these genes.

Auxin and Ethylene Enhance Flavonol Accumulation

To analyze changes in flavonol accumulation after treatment with IAA and ACC, we used a dye, diphenylboric acid 2-aminoethyl ester (DPBA), to visualize flavonols and their glycosylated derivatives in live seedling roots. DPBA fluorescence is absent in mutants that make no flavonols (Peer et al., 2001; Buer and Muday, 2004). DPBA fluoresces with distinct spectral properties in complex with kaempferol and quercetin (Neu, 1956; Sheahan and Rechnitz, 1992; Peer et al., 2001), with Q-DPBA having a longer wavelength fluorescence emission than K-DPBA. In Arabidopsis seedlings, DPBA fluorescence is largely due to binding to glycosylated flavonols, as aglycone flavonols are of low abundance or nondetectable in these tissues, as determined by HPLC analyses (Burbulis et al., 1996; Böttcher et al., 2008; Yonekura-Sakakibara et al., 2008). We optimized laser scanning confocal microscope (LSCM) settings to spectrally separate the fluorescence of K-DPBA and Q-DPBA (which reflect the fluorescence of the glycosylated forms of kaempferol and quercetin, respectively) by capturing a window of the emission spectrum for each compound where they do not overlap.

We detected the fluorescence spectra after excitation at 458 nm in vivo for Q-DPBA and K-DPBA using mutants and treatments with exogenous flavonols while maintaining consistent laser and gain settings, as shown in Figure 3. A K-DPBA spectrum was defined in the quercetin-deficient mutant, tt7-2 (Fig. 3B), and by treatment of tt4-2 with kaempferol (data not shown). In addition, DPBA fluorescence in tt4-2 fed with 10 μm quercetin provided a spectrum of quercetin fluorescence. Another report indicated that excitation with UV light enables visualization of isorhamnetin-DPBA fluorescence (Auger et al., 2010). We fed isorhamnetin to tt4 roots and observed at 458-nm excitation that isorhamnetin-DPBA fluorescence was barely detectable (Fig. 3A). Fluorescence-capture settings were tailored to these reference spectra and validated by the absence of fluorescence in untreated tt4-2 and the absence of Q-DPBA fluorescence in tt7-2 stained with DPBA (Fig. 3B) as well as the presence of a consistent pattern and ratio of Q-DPBA and K-DPBA fluorescence in wild-type roots.

Determination and validation of LSCM settings for kaempferol and quercetin measurement. A, Graph of the emission spectra for kaempferol (K), quercetin (Q), and isorhamnetin (IR) acquired with tt7-2 roots as well as isorhamnetin- and quercetin-fed tt4-2 roots, with the wavelength range used for K-DPBA collection shaded in green and the Q-DPBA collection window in gold. B, Confocal micrographs of 6-d-old DPBA-stained wild-type, tt4-2, tt7-2, and tt4-2 primary roots 6 h after treatment with 10 μm quercetin illustrate the specificity of these settings for detecting only K-DPBA in green or Q-DPBA in yellow. Bar = 100 μm.

Flavonol accumulation in roots grown on control medium and roots transferred for 8 h to medium containing 1 μm IAA or 1 μm ACC is shown in Figure 4A. This experiment was conducted 8 h after treatment, compared with the 4- to 6-h treatments used in the gene expression studies, as metabolite synthesis likely lags behind the gene expression events leading to enzyme synthesis. Root images were captured by LSCM with inset rectangles, which cover the same area and contain the same number of cells, defining the region used for the intensity measurements. Untreated wild-type roots exhibited K-DPBA staining throughout the meristematic and elongation zones, while Q-DPBA was more concentrated in the elongation zone. DPBA fluorescence appears to be both cytoplasmic and nuclear localized in these cells, which have not yet elongated and therefore contain several small provacuoles that are devoid of fluorescence. This subcellular localization pattern is consistent with previous reports that enzymes required for flavonoid synthesis and pathway products accumulate in the cytoplasm and nucleus (Saslowsky et al., 2005). IAA caused 2.5- and 3.1-fold increases in the amounts of K-DPBA and Q-DPBA fluorescence, respectively, as shown in Table II. We also examined DPBA fluorescence at a range of IAA concentrations and found robust increases that were proportional to the dose, over the range of 0.05, 0.1, 0.5, and 1 μm (data not shown). This increase in Q-DPBA signal was maximal in the transition between the elongation and differentiation zones, which is readily observed by the differences in cell elongation from the top to the bottom of the high-magnification images in Figure 4A. ACC caused a 3.2-fold increase in K-DPBA fluorescence and a 1.6-fold increase in Q-DPBA in a similar region of the root.

IAA and ACC increase flavonoid accumulation in primary roots and alter the distribution of these metabolites. A, Representative images of primary roots stained with DPBA 8 h after mock treatment or exposure to 1 μm IAA or 1 μm ACC. K-DPBA- and Q-DPBA-specific signals were collected, pseudocolored, and displayed as individual channels and an overlay. The right column shows higher magnification images that were used for quantification in the area defined by the red box, which contains seven cells. Mock-, IAA-, and ACC-treated roots are shown in rows one to three, respectively. Bar in left three columns = 100 μm; bar in right column = 50 μm. B, Comparison of DPBA and FIE-MS analyses of flavonoid accumulation in primary root tips in the elongation zone (DPBA) or in 5-mm root tips containing the elongation zone and an additional 4 mm (FIE-MS) after treatment with IAA and ACC. The two methods show a similar increase in flavonol accumulation 12 h after hormone treatment. # Significant difference between treated and untreated controls within a metabolite as determined by Student’s t test (P < 0.05). C, Six-day-old DPBA-stained wild-type, tir1-1, and ein2-5 primary roots. Representative images are from three separate analyses, each of which consisted of more than five micrographs each per genotype taken with identical gain and laser intensity. Gain settings were increased relative to A to better visualize DPBA staining in tir1-1 and ein2-5. Bar = 100 μm.

Table II.

DPBA fluorescence is altered in tir1-1, ein2-5, and myb12

We also verified that flavonol changes occur under identical conditions to those used for the gene expression studies and are not the result of transfer from one plate to another. DPBA staining was examined in roots grown on nylon filters, as used for the gene expression studies. Under those conditions, IAA and ACC enhanced DPBA fluorescence with a similar magnitude as in seedlings transferred to medium without filters (data not shown). We also examined DPBA staining in roots transferred from the growth plate to a control plate without added IAA or ACC. In these experiments, we found no change in DPBA fluorescence in response to transfer (data not shown). These results are consistent with a differential regulation of the pathway by IAA and ACC that creates specific metabolite profiles that are detected as changes in the ratio of Q-DPBA and K-DPBA fluorescence (Table II).

To directly measure the levels of specific flavonols in root tips, we used a flow injection electrospray mass spectrometry (FIE-MS) approach. Extracts from the apical 5 mm of single root tips containing the entire root elongation zone (which is localized within the first 1 mm) were analyzed by precursor ion scanning utilizing mass spectral parameters optimized for the detection of kaempferol, quercetin, and isorhamnetin (3′-methyl quercetin) and based on the production of the corresponding positively charged ions for the aglycone skeletons (287, 303, and 317 mass-to-charge ratio [m/_z_]), respectively. The mass values observed for the parent ions in combination with the fragment ions used for precursor ion scanning supported the assignment of the two major flavonol glycosides in Arabidopsis seedling roots as kaempferol-glucoside-rhamnoside and quercetin-glucoside-rhamnoside (Routaboul et al., 2006; Yonekura-Sakakibara et al., 2008). Eight additional flavonol glycosides were detected in the extracts as well, and these, in total, constituted less than 15% of the above two compounds. These were assigned as kaempferol-rhamnosyl(1→2)glucoside-rhamnoside, kaempferol-rhamnoside, kaempferol-glucoside, quercetin-rhamnosyl(1→2)glucoside-rhamnoside, quercetin-rhamnoside, quercetin-glucoside, isorhamnetin-rhamnoside-glucoside, and isorhamnetin-glucoside (data not shown). A comparison of results obtained by DPBA staining and FIE-MS analysis of the two must abundant kaempferol and quercetin glycosides 12 h after treatment with IAA and ACC versus an untreated control is shown in Figure 4B. Absolute values of the abundance of derivatives of the methylated form of quercetin, isorhamnetin, were also examined and are reported along with the unnormalized data for kaempferol and quercetin derivatives in Supplemental Figure S1. Isorhamnetin glycosides were detected at very low levels in untreated control root tips and increased in parallel with quercetin and kaempferol glycosides in response to IAA and ACC. However, isorhamnetin glycosides remained a minor component of the total flavonol content of the root tip and also did not fluoresce upon binding DPBA, as shown in Figure 3A. Although the FIE-MS technique showed a higher fold change (increase in relative peak area) in total flavonol accumulation and a less pronounced change in the quercetin-kaempferol ratio (Q/K ratio) after treatment with IAA or ACC than the quantification obtained using the DPBA staining technique, both trended in the same direction. The reported differences in the two methods likely reflect spatial differences, as the DPBA fluorescence was quantified in specific cells in the elongation zone while FIE-MS was performed with 5-mm root segments that include the elongation zone, the root tip, and an additional greater than 4-mm region of root beyond the elongation zone. This difference may also reflect the focus of the FIE analysis on the most abundant flavonol glycosides, rather than the total pool. These FIE-MS results thus validate the use of DPBA staining for flavonol quantitation and indicate that the observed changes in DPBA fluorescence after hormone treatment parallel changes in flavonol abundance.

Auxin and Ethylene Signaling Mutants Show Reduced Response to Either IAA or ACC

As the auxin and ethylene-insensitive mutants were found to have lower levels of accumulation of transcripts encoding enzymes of flavonol biosynthesis, we predicted that these genotypes would also exhibit reduced flavonol accumulation. To test this hypothesis, Q-DPBA and K-DPBA fluorescence was examined in primary roots of the wild type, tir1-1, and ein2-5, as shown in Figure 4C and quantified in Table II. To facilitate genotype and treatment comparisons, the DPBA fluorescence values in Table II are normalized to Q-DPBA values in untreated controls. Compared with the wild type, tir1-1 accumulated 60% less Q-DPBA and 40% less K-DPBA in the elongation zone (P < 0.001). The elongation zone of ein2-5 also showed a significant decrease in Q-DPBA (40%) and K-DPBA (50%) accumulation relative to the wild type (P < 0.005). The Q/K ratio was 2.5 in the wild type, 1.6 in tir1-1, and 3.0 in ein2-5. The lower Q/K ratio in tir1 is consistent with endogenous auxin signaling driving the expression of F3′H. ein2-5 exhibited reduced K-DPBA fluorescence at the root tip, whereas the distribution of DPBA staining in tir1-1 was similar to the wild type.

We also examined the flavonol-DPBA fluorescence in a mutant defective in MYB12. The untreated myb12 accumulated 30% and 39% less kaempferol and quercetin, respectively, than the untreated wild type (P < 0.03) but had a roughly equivalent Q/K ratio (Table II). Moreover, a mutation in _MYB12_ blocks IAA- and ACC-dependent flavonol accumulation increases with no significant change with either treatment, as compared with the untreated control (_P_ > 0.8). Interestingly, while mutations in TIR1, EIN2, and MYB12 all significantly reduce DPBA fluorescence (P < 0.05), each impacts the Q/K ratio in distinct ways.

In addition, we examined the effect of IAA and ACC added together on flavonol accumulation as judged by DPBA staining (Table II), and the effect was more additive than synergistic, as was the case for the induction of transcript levels by these hormones. The fold increase in both kaempferol and quercetin exceeded that of either individual treatment and was roughly equal to the sum of the individual treatments, with significant increases over either treatment with IAA or ACC alone (P < 0.01), further strengthening the case for independent signaling pathways for IAA- and ACC-mediated regulation of flavonoid metabolism. Additionally, in the tir1 and ein2 mutant backgrounds, we observed specific defects in IAA- and ACC-mediated flavonol accumulation, respectively, as well as opposite changes in the Q/K ratio. Conversely, mutations in MYB12 block both IAA- and ACC-mediated flavonol accumulation and have no significant effect on the Q/K ratio (Table II).

Auxin and Ethylene Induce Transcript and Flavonol Accumulation with Distinct Kinetics

The results above describe increases in the transcript abundance of genes encoding flavonol biosynthetic enzymes (4–6 h after treatment) and flavonol accumulation at 8 to 12 h, but with a varying effect between genes and metabolites. Yet, the timing for the gene expression and metabolite changes may be temporally distinct, and important changes may not be detected if only one time point is examined. Additionally, it is expected that expression changes in transcription factors should precede changes in target genes encoding pathway enzymes. Therefore, we performed time-course experiments to determine the kinetics of enhanced expression of an expanded set of genes encoding flavonoid biosynthetic enzymes in response to IAA and ACC. qRT-PCR was used to assay CHS, CHI, F3H, FLS, and F3′H transcript abundance at eight time points ranging from 0.5 to 24 h after treatment with 1 μm IAA and 1 μm ACC. Analysis of transcript abundance relative to an untreated control revealed transient increases in CHS, CHI, FLS, and F3′H, but not F3H, transcript levels after treatment with IAA, as shown in Figure 5A. CHS, CHI, and FLS transcripts reached a maximum around 2 h after treatment with a 3- to 4-fold increase and returned to baseline at 24 h. F3′H transcript levels increased 2.8-fold at 2 h after treatment but did not reach a maximum of 4.4-fold until 8 h, showing delayed kinetics compared with CHS.

In Arabidopsis roots, IAA and ACC differentially induce gene expression, which predicts metabolite accumulation kinetics. A and B, Kinetic analysis of CHS, F3′H, and FLS expression via qRT-PCR after treatment with 1 μm IAA (A) and ACC (B). Abundance of the mRNA from three genes relative to actin is represented as fold increase over untreated controls. C and D, Q-DPBA- and K-DPBA-specific fluorescence in primary roots stained with DPBA was monitored over time in response to IAA (C) and ACC (D). E and F, Kinetic analysis of IAA-induced (E) and ACC-induced (F) changes in CHS, MYB12, TTG1, and PAP1 expression determined from RNA samples used in A and B. Average and se of normalized gene expression from three pools of individuals are shown in A, B, E, and F; average and se of DPBA fluorescence from three independent trials containing eight individual roots each are shown in C and D.

A kinetic analysis of ACC-induced gene expression changes was also performed. A graph of transcript abundance over time after ACC treatment is shown in Figure 5B. CHS, CHI, and FLS increased 3- to 4-fold by 2 or 4 h after treatment and maintained that level until 8 h after treatment. The effects of ACC on F3′H transcript abundance were much more subtle, with no significant changes in transcript abundance observed until 4 h after treatment and a peak of only 1.7-fold at 8 h. The ACC-mediated increases in CHS, but not F3′H, transcript abundance predict that kaempferol accumulation should be more strongly induced by ACC than quercetin, consistent with the DPBA fluorescence shown in Figure 4.

The kinetics of accumulation of flavonols after treatment with 1 μm IAA or ACC are shown in Figure 5, C and D, respectively. After treatment with IAA, increases in K-DPBA fluorescence precede increases in Q-DPBA fluorescence but lag behind changes in CHS transcript abundance. In addition, F3′H transcript changes predict changes in quercetin accumulation, or more specifically, the ratio of F3′H to CHS transcript abundance seems to predict the ratio of quercetin to kaempferol accumulation (Fig. 5, A and C). Both kaempferol and quercetin remained significantly above control levels 24 h after treatment with IAA, even though the enhanced gene expression had returned close to untreated levels. This is consistent with enzyme levels remaining elevated and the resultant increase in metabolism persisting after transcripts have returned to baseline levels.

The kinetics of the accumulation of flavonols in response to ACC differ from the IAA kinetics in profile and in maximal response. K-DPBA increased much more substantially than Q-DPBA and at earlier time points than in response to IAA treatment. The slower rate of quercetin accumulation caused by ACC relative to IAA was mirrored by the less potent effect of ACC on F3′H transcript abundance. These results indicate that IAA and ACC enhance carbon flow through the flavonoid pathway, but with different effects on the abundance of quercetin and kaempferol, suggesting that metabolic flux may be differentially regulated by IAA and ACC through the regulation of F3′H transcript and presumably enzyme levels.

IAA and ACC Increase CHSpro:CHS-GFP Fluorescence and CHS Transcripts with Similar Kinetics

We also tested the hypothesis that treatments that increased CHS mRNA accumulation would have the same effect on CHS protein levels and asked if these changes occurred in the same regions of the root where flavonol accumulation was observed. Plants harboring a CHSpro:CHS-GFP translational fusion, expressed in the tt4-11 background, were imaged after treatment with IAA and ACC as shown in Figure 6A. Both 1 μm IAA and 1 μm ACC caused GFP fluorescence to increase in the distal and central elongation zones of primary roots. CHSpro:CHS-GFP fluorescence was detected in all radial cell layers, with slightly lower levels in the epidermis than internal tissues, which are difficult to visualize without the cortical fluorescence signal becoming saturated.

CHSpro:CHS-GFP is induced in the elongation zone of primary roots in response to IAA and ACC. A, Representative images of CHSpro:CHS-GFP in primary roots of 6-d-old seedlings counterstained with propidium iodide (shown in white) at 8 h after the indicated treatment. A digitally reconstructed maximum projection is shown from a z-stack of micrographs, as is a cross-section showing a single optical slice from the center of the root. Representative images are shown from two trials in which 20 total micrographs were taken. Bar = 100 μm. B, Comparison of CHSpro:CHS-GFP fluorescence, CHS transcript, and flavonol accumulation during a time course after IAA treatment. C, Comparison of CHSpro:CHS-GFP fluorescence, CHS transcript, and flavonol accumulation during a time course after ACC treatment. For CHSpro:CHS-GFP, the average and se are shown from two separate experiments, quantitatively imaging six roots per time point. The transcript data from Figure 5 is included for comparison. To reflect the total flavonol pool, the Q-DPBA and K-DPBA fluorescence from Figure 5 are combined.

A comparison of CHSpro:CHS-GFP fluorescence, CHS mRNA levels, and total flavonol accumulation after treatment with IAA and ACC is shown in Figure 6, B and C. GFP fluorescence increased in response to IAA and ACC treatment, but with slower kinetics than the increase in CHS transcript abundance. To compare the timing of increases in flavonol abundance, CHS mRNA, and CHS-GFP fluorescence, the levels of Q-DPBA and K-DPBA were summed to reflect the total flavonol pool. In both IAA and ACC treatment, flavonol accumulation was proportional to CHSpro:CHS-GFP fluorescence. The increase in CHS transcript abundance preceded increases in protein abundance, which was then followed by changes in flavonoid accumulation. The elevated flavonol levels are sustained even when the RNA and GFP signals are decreased, consistent with greater stability and longevity of these metabolites.

Expression of Genes Encoding Transcriptional Regulators Linked to the Phenylpropanoid Pathway Is Also Induced by IAA and ACC

We examined gene expression in response to auxin and ethylene treatment in publicly available and previously published microarray data sets (Armstrong et al., 2004; Redman et al., 2004; Vanneste et al., 2005) and looked for transcription factors that might be induced by hormones that coordinate the expression of flavonoid genes in response to both auxin and ethylene as well as the expression of transcription factors previously linked to the regulation of flavonoid structural genes. The increased accumulation of MYB12, TTG1, and PAP1 to IAA and ACC in these microarray data sets is shown in Supplemental Table S1.

Therefore, we used qRT-PCR to examine the levels of MYB12, PAP1, and TTG1 transcripts after treatment with 1 μm IAA and 1 μm ACC, as shown in Table I and Figure 5. The abundance of all four transcripts increased after treatment with IAA and ACC. The large magnitude response of MYB12 at 6 h after treatment (14-fold) exceeds the changes in the transcripts for pathway enzymes described above and contrasts with the comparatively modest increases of PAP1 and TTG1 (Table I). The greater magnitude changes in MYB12 may be due to the relatively low transcript abundance of this gene in untreated samples (Table I) compared with the genes encoding pathway enzymes.

We also examined the levels of transcripts encoding MYB12, TTG1, and PAP1 in the tir1-1 and ein2-5 mutant backgrounds to ask if the changes in transcript abundance for these genes required these auxin and ethylene signaling proteins, as shown in Figure 7A. The large increase in all three transcripts after auxin treatment was lost in tir1-1, and the ACC response was lost in ein2-5. In both cases, the transcript abundance of all three genes was not significantly different from untreated controls within the genotype (P > 0.3). Therefore, it is clear that the hormonally induced increase in these transcription factors requires intact TIR1 and EIN2 and that these transcript changes are also modulated through independent pathways.

Auxin and ethylene responsiveness of PAP1 and TTG1 in roots requires TIR1, EIN2, and MYB12. A, The basal level and fold induction of MYB12, PAP1, and TTG1 transcript abundance 6 h after treatment with IAA and ACC is shown in all genotypes. TIR1 and EIN2 are necessary for auxin- and ethylene-dependent gene expression changes, respectively. All data are normalized to untreated wild-type (WT) expression levels. B, Mutations in MYB12 prevent the induction of PAP1 and TTG1 transcript changes after IAA and ACC treatment. Mean and se of three biological replicates are shown. * Significant difference (P < 0.05) between the mutant and the wild type within a treatment; # significant difference between treated and untreated controls within a genotype as determined by Student’s t test (P < 0.05).

As MYB12 has been shown to be required for flavonol synthesis in Arabidopsis roots and the myb12 mutant line did not exhibit hormone-induced increases in DPBA fluorescence (Table II), we tested whether IAA and ACC treatments would alter CHS, F3′H, and FLS transcript abundance in the myb12 mutant, as shown in Figure 2. In untreated seedlings, the levels of these flavonoid gene transcripts were lower than in the wild type, and IAA and ACC did not cause changes in transcript levels relative to untreated controls. These results are consistent with a requirement for MYB12 for the expression of these flavonoid genes, both in controls and in response to IAA and ACC treatments. An analysis of PAP1 and TTG1 mRNA levels after treatment with IAA and ACC in the myb12 mutant is shown in Figure 7B. There are significant decreases in the abundance of transcript encoded by these genes in the untreated mutant relative to the wild type (P < 0.05). In addition, treatment with IAA and ACC caused no significant change in these genes in myb12 (Supplemental Fig. 3). These data suggest a consensus role for MYB12 in regulating the auxin and ethylene responsiveness of both flavonol pathway genes and the transcription regulators that have been linked to the control of phenylpropanoid metabolism.

IAA- and ACC-Enhanced Accumulation of Transcripts Encoding Transcriptional Regulators Precedes the Accumulation of Transcripts of Pathway Enzymes

We examined the transcript abundance of MYB12, TTG1, and PAP1 after IAA and ACC treatment and asked if the kinetics were consistent with these transcriptional regulators increasing prior to the induction of pathway genes, as shown in Figure 5, E and F. Increased abundance of transcripts encoding these transcription factors was initially detectable 0.5 h after treatment, much sooner than for CHS, CHI, F3′H, or FLS. The large magnitude response of MYB12 contrasts with the comparatively modest increases observed for PAP1 and TTG1. The PAP1 and TTG1 changes, which are partially masked in the complete time course graphs, are highlighted in an inset showing the first 4 h of the response (Fig. 5) as well as quantified in Supplemental Table S2 at 0.5 and 2 h after treatment. In both cases, the peak in transcript accumulation of the MYB transcription factors (MYB12, PAP1 [_MYB75_]) preceded the peak in the transcripts of the non-MYB transcriptional regulator (TTG1). These data are consistent with MYB12 acting upstream of these other proteins, as are the myb12 mutant data shown in Figure 7A. These results suggest that the flavonol biosynthetic genes are independently regulated by auxin and ethylene, with an absolute requirement for MYB12 for expression under all conditions. We examined the regulatory regions of these key genes and searched for consensus elements for binding of transcription factors. We searched for auxin response element (AuxRE) sites, which bind ARF, using the consensus AuxRE sequence TGTCTC (Ulmasov et al., 1995), ERFs using the consensus ethylene response element (ERE) sequence TAAGAGCCGCC (Ohme-Takagi and Shinshi, 1995), and the MYB RE sequence AACCTACC (Hartmann et al., 1998). We identified potential sites for binding of these transcription factors in many of these promoter regions, as shown in Supplemental Table S3. Putative AuxRE sites were found in all these genes, while sequences that resemble the ERE site were also found, with the weakest similarity found in F3′H, consistent with the weak induction of F3′H by ACC. All of these genes have well-conserved MYB response elements, with the weakest similarity found in the F3H regulatory region, which showed neither IAA- nor ACC-dependent transcript changes.

Basipetal Auxin Transport and Dependent Processes Are Altered Similarly in tt4-2 and tt7-2

Previous experiments have shown elevated IAA transport in tt4-2 mutants that make no flavonols, consistent with the loss of a negative regulator (Brown et al., 2001; Buer and Muday, 2004). The results presented above indicate that IAA and ACC lead to different levels of kaempferol and quercetin in the root. We asked whether kaempferol and/or quercetin are the negative regulators of basipetal, or shoot-ward, IAA transport. As tt4-2 accumulates no flavonols and tt7-2 produces kaempferol but not quercetin, comparing auxin transport in these two mutants can elucidate the role of kaempferol and quercetin in auxin transport inhibition. Both tt4-2 and tt7-2 exhibited statistically significant increases in basipetal auxin transport relative to the wild type (P < 0.01), as shown in Figure 8A. These data suggest that quercetin, but not kaempferol, is an inhibitor of root basipetal auxin transport. Treatment with 1 μm ACC increased auxin transport capacity in the wild type, tt4-2, and tt7-2, with 2- to 2.5-fold increases in all three genotypes, consistent with a mechanism by which ACC positively regulates transport that is independent of flavonols. To ask if auxin transport-dependent processes are also regulated by quercetin, but not by kaempferol, the gravitropic response was measured in wild-type, tt4-2, and tt7-2 plants treated with and without 1 μm ACC. A high-resolution morphometric analysis of root tip angle after reorientation is shown in Figure 8B. Both tt4-2 and tt7-2 exhibited an impaired gravitropic response, as compared with the wild type. In the wild type, ACC leads to a substantial reduction in gravitropism, which is of greater magnitude than the effect of tt4-2 and tt7-2 mutations. In both tt4-2 and tt7-2, ACC treatment resulted in no inhibition of gravitropism. In Figure 8, the slight increase in gravitropic response in tt4-2 and tt7-2 after ACC treatment is not statistically significant. The identical phenotypes of tt4-2 and tt7-2, along with the similar lack of ACC response in the two lines, suggest that quercetin, but not kaempferol, regulates the gravitropic response.

tt4-2 and tt7-2 exhibit altered auxin transport and signaling and dependent processes. A, Basipetal [3H]IAA transport in the wild type (wt), tt4-2, and tt7-2 is shown. Average and se of three independent trials of eight individuals are shown. B, Morphometric analysis was used to quantify the tip angle of roots after gravitropic stimulation. The wild type, tt4-2, and tt7-2 in the presence and absence of 1 μm ACC are shown (mean and se of more than seven individual roots). Data from tt4-2 and tt7-2 as well as these genotypes treated with ACC are overlapping, and in some cases, difficult to visualize. C, Growth inhibition by 1 μm ACC in the wild type, tt4-2, and tt7-2. The tt4-2 growth responses in both the presence and absence of ACC are masked by overlapping tt7-2 growth responses. Mean and se are shown for three independent trials, each containing six individuals. D, Comparison of the wild type (WT) and tt4-2 expressing the DR5:vYFP auxin-responsive promoter 12 h after treatment. tt4-2 exhibits reduced ACC induction of DR5 expression, especially in the cortical cell layer (white boxes) and in the lateral root cap (arrows), where the ACC effect on DR5 activation is most apparent. Representative images from two independent trials of eight individuals are shown. Bar = 100 μm.

The inhibition of primary root elongation by ACC was also analyzed in the wild type, tt4-2, and tt7-2 (Fig. 8C). The tt4-2 and tt7-2 mutants exhibit identical growth kinetics, which completely overlay on this graph, and which are slightly faster than for the wild type. The wild type responded to 1 μm ACC with a profound reduction in growth. In the presence of ACC, both mutants have identical growth kinetics, which indicates identical reductions in root elongation responses to exogenous ACC in both mutants. The similar phenotypes of tt4-2 and tt7-2 indicate that quercetin, but not kaempferol, is required for the maximal inhibitory effect of exogenous ACC on root elongation rate. The timing of the first divergence in growth between the wild type and tt4-2 or tt7-2 at 8 h is concurrent with the plateau of flavonoid accumulation shown in Figure 5. Thus, elongation growth and gravitropism phenotypes of tt4-2 and tt7-2 in response to ACC become more apparent after observed flavonoid accumulation increases, consistent with a role for quercetin in modulating the inhibition of growth and gravity response by ACC treatment.

Induction of Auxin Signaling in the Elongation Zone of Roots by ACC Is Reduced in tt4-2

To probe the physiological basis of ACC-mediated growth inhibition and its dependence on flavonols, the induction of auxin signaling in the elongation zone of tt4-2 was analyzed and compared with the wild type using the nucleus-localized DR5:vYFP auxin-responsive reporter (Laskowski et al., 2008). Representative images of eight to 12 individuals each of the wild type and tt4-2 expressing DR5:vYFP 12 h after treatment with 1 μm ACC are shown in Figure 8D. These images were captured with identical gain, intensity, and pinhole settings to allow quantitative comparison between seedlings. White boxes are drawn to highlight the cortical cells in the elongation zone, where the ACC-induced auxin signaling should negatively influence elongation growth. ACC causes auxin signaling increases in wild-type lateral root cap cells, which is highlighted by the arrows. These changes were consistently observed in all wild-type roots and were dramatically reduced in all tt4-2 roots that were examined, as shown in the representative in Figure 8D. The increase in auxin signaling in the central cylinder, epidermis, cortex, and root cap of the elongation zone in response to ACC in the wild type was consistent with previous reports using DR5:GUS (Růzicka et al., 2007). The reduction in this auxin response in tt4-2 is consistent with flavonols affecting the ACC response by altering auxin distribution. These results demonstrate that products of the flavonoid pathway are required for the induction of DR5 in the cortex of the elongation zone of primary roots in response to 1 μm ACC.

DISCUSSION

Auxin and ethylene regulate plant growth and development through tightly regulated and well-studied transcriptional networks (Kendrick and Chang, 2008; Chapman and Estelle, 2009; Stepanova and Alonso, 2009). In recent years, cross talk between auxin and ethylene has been shown to regulate root elongation (Růzicka et al., 2007; Stepanova et al., 2007; Swarup et al., 2007), gravitropism (Rahman et al., 2001; Buer et al., 2006; Muday et al., 2006), lateral root formation (Ivanchenko et al., 2008; Negi et al., 2008, 2010), and hypocotyl growth, including elongation, hook opening, and gravitropism (Vandenbussche et al., 2003, 2005; Muday et al., 2006). These studies have provided evidence that auxin regulates ethylene synthesis and that ethylene regulates auxin synthesis and transport (Stepanova and Alonso, 2009). Several studies have looked at the global transcriptional response to either exogenously applied or endogenously overproduced auxin or ethylene (Zhong and Burns, 2003; Redman et al., 2004; Vanneste et al., 2005; Paponov et al., 2008). Only one study in Arabidopsis examined the global transcript accumulation changes in response to IAA and ACC within the same experiment and identified transcripts that change by common or distinct pathways (Stepanova et al., 2007). These results suggest that there are target genes that are regulated by both hormones through distinct signaling pathways, yet the functional significance of many of these transcriptional changes in controlling growth and development has not been determined.

Therefore, we examined the effects of auxin and ethylene on the expression of genes encoding enzymes of the flavonoid biosynthetic pathway, as previous results have suggested that metabolites produced by this pathway are elevated in response to treatments with these two hormones (Buer and Muday, 2004; Buer et al., 2006), which in turn regulate plant growth and development (Brown et al., 2001; Buer and Muday, 2004; Peer et al., 2004; Buer and Djordjevic, 2009). Our experiments showed that CHS, CHI, and FLS transcript abundance was increased by both IAA and ACC, while F3′H transcription was significantly increased by IAA alone, and F3H and DFR were unchanged. Kinetic analyses further showed that mRNAs encoding the transcriptional regulators MYB12, PAP1, and TTG1 preceded the induction of CHS, CHI, FLS, and F3′H mRNA levels, with changes in MYB12 transcript levels by far the most dramatic and earliest, suggesting that the expression not only of flavonoid genes but also of PAP1 and TTG1 may be dependent upon MYB12 activity. As expected, the changes in flavonoid gene expression preceded increases in CHSpro:CHS-GFP and flavonol levels in response to hormone treatment. This sequential increase in transcripts of a regulatory module of transcription factors in advance of proteins needed for cellular responses is now the subject of studies focused on unraveling the higher order temporal coordinating networks that regulate these events (Yosef and Regev, 2011).

These kinetic analyses also suggest that hormone treatment changes the relative abundance of specific metabolites by regulating the expression of the branch point enzyme F3′H, which is required for quercetin synthesis (Fig. 1). The results suggest a model in which low abundance of F3′H relative to other pathway enzymes, together with delayed induction of F3′H expression in response to IAA, results in a transient peak in kaempferol accumulation followed by a later increase in quercetin accumulation. The correlation of the Q/K ratio with the F3′H/CHS transcript ratio suggests that metabolic partitioning of intermediates in response to hormone treatment may be regulated via transcriptional control of enzyme synthesis. The different peak expression times of CHS and F3′H are consistent with higher order temporal coordination (Yosef and Regev, 2011), in which coordinated timing motifs are found at metabolic branch points and where sequential activation of genes encoding pathway enzymes follow the order of steps in that pathway (Zaslaver et al., 2004).

Several well-characterized auxin and ethylene signaling mutants were used to explore the mechanisms by which IAA and ACC induce flavonoid gene expression and flavonol accumulation in seedling roots. TIR1 encodes an auxin receptor, which is essential for auxin-induced gene expression (Dharmasiri et al., 2005), while ETR1 and EIN2 function as ethylene receptor and signaling proteins, respectively (Bleecker et al., 1988; Chang et al., 1993; Alonso et al., 1999). In these signaling mutants, lower levels of transcripts and flavonols were found than in the wild type and, as anticipated, tir1-1 did not exhibit enhanced transcript abundance in the presence of IAA, while ein2-5 and etr1-3 did not respond to ACC treatment. The nearly complete absence of transcriptional responses in tir1 and etr1 is somewhat surprising, as these mutants exhibit reduced, but not absent, growth response to high concentrations of auxin and ethylene, respectively (Ruegger et al., 1998; Hall et al., 1999). However, this experiment was performed at a dose of ACC that does not affect root elongation or branching in etr1-3 (Negi et al., 2008) and is equivalent to a dose of ethylene gas that also does not affect root elongation in the etr1-3 mutant (Hall et al., 1999; Larsen and Chang, 2001).

A particularly striking result is the normal response of ein2-5 and etr1-3 to IAA and tir1-1 to ACC with regard to both flavonoid gene expression (Fig. 2) and flavonol accumulation (Table II). This finding, together with the additive effects of IAA and ACC on CHS transcript and flavonol levels in wild-type seedlings, suggests that these two hormones act through independent signaling pathways. Conversely, mutations in MYB12 abolish the induction of CHS, FLS, and F3′H transcripts and flavonol levels by either IAA or ACC, indicating that this transcription factor is required for both responses. Our results add new information beyond previous studies showing that, although F3′H expression is elevated in a MYB12 overexpressor (Mehrtens et al., 2005), expression of F3H, but not F3′H and DFR, is regulated by MYB12 in cell cultures and whole seedlings. This suggests the possibility that a higher level of MYB12 may be required to induce F3′H expression and could explain the delayed induction of this gene, and the attendant shift in the kaempferol-to-quercetin ratio, in response to IAA and ACC treatment.

The model presented in Figure 9 summarizes our results and suggests a hierarchical requirement for transcriptional regulators, in which the well-characterized and primary auxin and ethylene signaling mechanisms converge on MYB12 to induce flavonoid gene expression and, subsequently, the metabolite products. Consistent with this model, the promoter regions of CHS, CHI, FLS, and MYB12 contain putative AuxRE and ERE (Ohme-Takagi and Shinshi, 1995; Ulmasov et al., 1995; Hartmann et al., 1998), while F3′H contains a highly conserved AuxRE but no well-conserved ERE. This is consistent with the absence of changes in F3′H transcript levels after ACC treatment. These experiments do not resolve whether MYB12 interacts with the auxin and ethylene transcriptional machinery. One recent study has suggested the presence of physical interactions between MYB77 and ARF7 that regulate a number of auxin-responsive genes (Shin et al., 2007), suggesting the possibility that MYB12 might work in complex with an ARF to induce the expression of flavonoid pathway genes.

Order of signaling events in hormone-induced flavonol metabolism. A combination of auxin- and ethylene-specific and MYB-dependent transcriptional control may explain the observed data. TIR1- and ARF-dependent auxin signaling activates the auxin-responsive elements of the pathway genes shown, which include F3′H. In parallel, MYB12 is activated, leading to further up-regulation of the same pathway genes and PAP1 and TTG1 through their MYB-responsive elements. The line between MYB12 and PAP1 and TTG1 is dotted based on the complexity of interpreting these mutant data, as described in the text. EIN2- and ETR1-dependent ethylene signaling also increases the expression of a subset of pathway genes, excluding F3′H, through the activation of ethylene-responsive elements. This auxin-specific regulation of F3′H may explain the observed differences in the Q/K ratio after treatment with IAA and ACC. [See online article for color version of this figure.]

Additionally, in myb12, the elevated PAP1 and TTG1 transcript levels after IAA and ACC are lost (Fig. 7B; Supplemental Fig. S3), suggesting an essential and upstream role for this MYB transcription factor in these auxin- and ethylene-driven transcriptional changes. As the myb12-1f allele that we used is predicted to encode a truncated MYB12 protein, an alternative possibility is that synthesis of this defective protein may interfere with the expression of PAP1 and TTG1. This possibility is represented in Figure 9 by a dashed arrow between MYB12 and PAP1/TTG1. One interesting surprise from these results is that, although PAP1 and TTG1 transcript accumulation increases after IAA and ACC treatment, transcript of a potential target of these transcriptional regulators, DFR, does not increase. This contrasts with the fact that anthocyanins accumulate in roots in the original activation-tagged pap1 line (Borevitz et al., 2000); however, this may also reflect substantially higher levels of PAP1 in this line and/or the transport of anthocyanins from aboveground tissues. We hypothesize that the lack of detectable DFR transcript is due to our analysis of RNA isolated from only root tissue, in contrast to previous studies on whole seedlings or leaf tissues (Tohge et al., 2005; Cominelli et al., 2008). This is supported by our ability to detect DFR in root genomic DNA but not in root-derived cDNA. There may be tissue-specific transcription factors that are not expressed in roots that work with PAP1 and TTG1 and that may be required for the activation of F3H and DFR. Demonstration of the precise transcription factor complexes that regulate the expression of these genes is beyond the scope of this work, but the precise interplay between MYB and auxin/ethylene-specific signaling pathways that control flavonol synthesis is an exciting avenue of future study.

In addition to the insights provided by high-temporal-resolution analyses, examination of transcript and metabolite accumulation with high spatial resolution is essential for linking these responses to the root growth and developmental responses controlled by these metabolites. Visualization of CHSpro:CHS-GFP fluorescence and DPBA-stained flavonols along the longitudinal axis of the root indicates that the elongation zone is the site of maximal flavonol production, consistent with previous results involving immunolocalization of CHS and CHI and DPBA staining of flavonoid end products (Saslowsky and Winkel-Shirley, 2001; Saslowsky et al., 2005). Our studies did not examine the localization of the transcripts encoding pathway enzymes, but previously published microarray data (Brady et al., 2007) that examined transcript accumulation with high spatial resolution show that transcripts encoding core enzymes in flavonoid synthesis are all similarly distributed, with high expression in the central elongation and differentiation zones (Supplemental Fig. 2). This indicates that the accumulation of flavonoids at this site is the result of localized induction of flavonoid gene expression.

These experiments were all performed with seedlings grown on transparent agar plates, as this is standard for experiments on root development, hormone signaling, and gene expression (Brady et al., 2007; Růzicka et al., 2007; Stepanova et al., 2007; Swarup et al., 2007; Péret et al., 2009). There is conflicting evidence on whether flavonoids also accumulate in roots when shoots are grown in the light and roots are grown in the dark. One report detected flavonols by in situ DPBA staining of plants grown with roots in the dark and shoots in the light (Buer and Muday, 2004), while another did not find flavonols in roots grown in darkness and examined in extracts separated by high-pressure thin-layer chromatography followed by DPBA staining (Stracke et al., 2010a). Accumulation of flavonols in dark-grown roots may be consistent with long-distance communication of light-dependent signals to the roots or movement of flavonols from the shoot into the root (Buer et al., 2007, 2008, 2010). The nature of this signal may in fact be flavonoid precursors themselves, which have been shown to move from an application site on the cotyledons to the root tip (Buer et al., 2007) or from wild-type shoots across a graft junction into tt4 roots (Buer et al., 2008). It is attractive to speculate that as Arabidopsis roots grow close to the soil surface, light-induced flavonoid synthesis and the positive effect of flavonoids on gravitropism may contribute to guiding roots deeper into the soil for maximal productivity.

The location of the transcripts, flavonoid biosynthetic enzymes that they encode, and flavonoid metabolites in the tissues that exhibit basipetal (or shoot-ward) auxin transport provides an ideal situation to test the role of specific flavonols on auxin transport and dependent processes. The epidermal, cortical, and lateral root cap cell layers in the root express auxin transport proteins essential for both basipetal IAA transport and gravitropism, including PIN2 and AUX1 (Muday and Rahman, 2006). Differential accumulation of flavonoid intermediates after treatment with IAA or ACC suggests that the flavonols, kaempferol and quercetin, and their glycosylated derivatives may have unique nonoverlapping functions. Analysis of basipetal or shoot-ward IAA transport, gravitropism, and primary root elongation in wild-type, tt4-2, and tt7-2 seedlings in the presence and absence of 1 μm ACC determined that tt4-2 and tt7-2 have identical auxin transport capacity, reductions in gravitropism, and responses to ACC treatment. This finding suggests that the absence of quercetin and its glycosylated and methylated derivatives, which include isorhamnetin, may be the cause of these phenotypes in both mutants. Additionally, flavonols are necessary for the full induction of auxin-induced gene expression in the outer cell layers of the elongation zone. This result, coupled with the partial insensitivity of tt4-2 to the negative effects of ACC treatment on root elongation, suggests that induction of the flavonoid pathway by ACC may be a mechanism by which the observed increases in auxin signaling negatively influence elongation growth.

Consistent with a role for quercetin in regulating basipetal IAA transport and gravitropism, quercetin treatment of the pin2 mutant, which has a defect in an IAA efflux protein that mediates basipetal IAA transport (Chen et al., 1998), restores the gravitropic response and the formation of gradients of auxin-induced gene expression across gravity-stimulated roots (Santelia et al., 2008). Our results contrast with a previous study that did not detect elevated IAA transport from the shoot apex into the root in the tt7-2 mutant but did detect elevated transport in tt4-2 (Peer et al., 2004). For the acropetal (or root-ward) IAA transport stream, these results suggested that kaempferol was sufficient for the regulation of auxin transport (Peer et al., 2004). We examined a different polar auxin transport stream in a different tissue, using a different tt7-2 allele, which was grown under different media conditions. Therefore, it is not yet clear whether quercetin (and its glycosylated and methylated derivatives) are the only flavonols actively regulating auxin transport beyond the basipetal IAA transport stream in roots. Interestingly, preliminary analyses of acropetal transport with the alleles described here showed that the tt4-2 and tt7-2 phenotypes are not equivalent, with tt7-2 showing an intermediate phenotype between the wild type and tt4-2 (data not shown). Furthermore, kaempferol and its derivatives have been shown to have distinct developmental roles in leaves based on analysis of the rol1-3 mutant, which is defective in Rha biosynthesis and flavonoid glycosylation (Ringli et al., 2008). This suggests that, in other tissues and auxin transport streams, kaempferol and its derivatives may have a distinct function.

The finding that treatment with ACC increased auxin transport capacity in the wild type, tt4-2, and tt7-2 is consistent with limited changes in the quercetin levels by this treatment and suggested a flavonoid-independent mechanism for enhanced IAA transport. The auxin transport proteins, PIN2 and AUX1, have also been shown to be expressed at higher levels at the root tip after ACC treatment (Růzicka et al., 2007), which may lead to this elevated root basipetal IAA transport. Similarly, ACC increases root acropetal IAA transport along with elevated PIN3 and PIN7 transcripts and fluorescent protein constructs, and the presence of functional PIN3 and PIN7 protein is required for this enhancement (D.R. Lewis, S. Negi, and G.K. Muday, unpublished data). Therefore, enhanced IAA transport in response to ACC transport seems to act through PIN proteins, while flavonoids likely target another class of auxin transporters, the ABCB family (Geisler et al., 2005).

CONCLUSION

In this study, we investigated the hormonal control of the flavonol biosynthetic pathway by measuring pathway enzyme gene expression, protein accumulation, and metabolite levels after treatment with the auxin, IAA, or the ethylene precursor, ACC. Mutant analysis revealed that the signaling pathways for auxin- and ethylene-mediated flavonol increases are distinct but intersect at the known positive regulator of flavonoid metabolism, MYB12. Transcriptional effects of hormone treatment on genes encoding pathway enzyme genes are rapid and transient and proceeded by increases in transcripts encoding transcriptional regulators of flavonoid metabolism, while the accumulation of metabolites is prolonged. Auxin- and ethylene-induced increases in flavonol accumulation as measured by DPBA fluorescence was most apparent in the elongation zone, well positioned to contribute to root elongation and gravitropism, and in the same tissues in which enzymes of flavonol synthesis and the transcripts that encode them have been localized. Furthermore, treatment with these two hormones yielded a different F3′H/CHS gene expression ratio, which then leads to distinct quercetin/kaempferol metabolite accumulation, consistent with the unique functional activities of these two flavonols. The tt4 mutant, which makes no flavonols, and the tt7 mutant, which makes kaempferol but not quercetin, exhibited equivalent phenotypes in basipetal auxin transport, elongation growth, and gravitropism, indicating that quercetin is the active modulator of auxin transport in this specific transport stream. Together, these data suggest the existence of distinct hormonal controls of this important biosynthetic pathway, the products of which in turn regulate plant growth and development, including the regulation of auxin transport and dependent growth processes, such as root gravitropism, elongation, and lateral root development.

MATERIALS AND METHODS

Plant Growth

Arabidopsis (Arabidopsis thaliana) plants were grown on 1× Murashige and Skoog medium (Caisson Labs), pH 5.6, Murashige and Skoog vitamins, and 0.8% agar, buffered with 0.05% MES (Sigma) and supplemented with 1.5% Suc. After stratification for 48 h at 4°C, plates were removed to vertical racks and placed under 100 μmol m−2 s−2 cool-white light. All assays were conducted at 6 d from transfer to the light, which is generally 4 to 5 d after germination, and at equivalent times of day to control for any circadian effects on measured processes. Lines used were tir1-1 (Ruegger et al., 1998), seeds generously provided by Mark Estelle; and etr1-3, ein2-5, tt4-2, and tt4-11 (SALK 020583), as described previously (Buer et al., 2006). The tt4-2 mutant is a backcross of tt4(2YY6) to remove an unlinked max4 mutation (Buer and Muday, 2004; Bennett et al., 2006). The tt7-2 (SALK 053394), tt4-11 (SALK 020583), and _myb12_-1f (stock no. CS9602) plants were generously provided by the Arabidopsis Biological Resource Center. The tt7-2 mutant was shown to be homozygous for insertion in the second exon using PCR (M.V. Ramirez and B.S.J. Winkel, unpublished data).

IAA and ACC Treatments and RNA Isolation

RNA was isolated from seedlings grown on a nylon screen (03-100/32; Sefar Filtration) as described previously (Levesque et al., 2006). Plants were germinated on a screen pressed tightly against control medium, with approximately 100 seedlings per plate, then on the 6th d after transfer to the light, the filter was transferred to a control plate or to medium supplemented with 1 μm IAA or 1 μm ACC for the indicated times. At the end of the treatment period, a razor blade was used to align the roots by dragging the plants to the center line and top of the plate, where the roots were excised, carefully excluding the root-shoot junction. Samples were promptly frozen in liquid nitrogen and stored at −80°C until RNA isolation.

Samples were ground using a liquid nitrogen-chilled plastic pestle and a power drill for 5 s, and Trizol reagent (Invitrogen) or lysis buffer (Qiagen) was added immediately. The remainder of the RNA isolation was performed according to the supplied protocol for isolating RNA from the lysate by either phenol chloroform phase separation followed by ethanol precipitation using the Trizol system or binding, wash, and elution from a column with the Qiagen Plant RNeasy kit. Nucleic acids were resuspended/eluted in 30 μL of 10 mm Tris-HCl in diethyl pyrocarbonate-treated water and digested with DNase (Promega) for 30 min at 37°C. RNA samples were quantified by _A_260 using a nanodrop spectrophotometer (Nanodrop Technologies). RNA concentrations were standardized to 150 ng μL−1 ± 10% by the addition of 10 mm Tri-HCl, pH 8.0. Each sample yielded approximately 4.5 μg of RNA.

cDNA Synthesis and qRT-PCR Analysis

Samples containing 900 ng of RNA were preincubated with a 1:1 mixture of oligo(dT) and random hexamer primers, then combined with a cDNA synthesis master mix containing SuperScript III enzyme (Invitrogen) The resultant cDNA/RNA mixture was stored at −20°C. After digestion with RNase (Invitrogen), the _A_260 was measured using a nanodrop spectrophotometer (Thermo Scientific) to ensure equal efficiency in the cDNA synthesis reactions between samples. qRT-PCR analysis of reverse-transcribed RNA was performed on an Applied Biosystems 7600-fast thermal cycler using SYBR Green detection chemistry.

Primers for CHS, F3′H, FLS, and F3H were designed with Primer Express software (Applied Biosystems) or Beacon Designer 7 (Premier Biosoft) using default parameters. Primers for PAP1, TTG1, CHI, and MYB12 were designed with the primer3 primer design software (Rozen and Skaletsky, 2000). These primers are listed in Supplemental Table S4. The efficiency coefficient E (Pfaffl, 2001) was calculated for all primer sets individually by plotting the relationship between cycle threshold (Ct) value and log[cDNA]. A standard curve was prepared, and for all primer sets used, we verified that the relationship was linear over a 1,000-fold concentration range, using the standard curve method (Applied Biosystems Technical Bulletin No. 2); the resultant primer efficiencies were used in an efficiency-corrected ΔΔCt formula. E values were as follows: ACTIN2 = 1.9, CHS = 2.0, F3′H = 2.1, FLS = 1.9, MYB12 = 3.0, CHI = 2.0, TTG1 = 2.0, and PAP1 = 2.0. The unusually high E value of the MYB12 primers represents a less efficient detection of changes in cDNA concentration, although over the entire concentration range tested, the relationship between log[cDNA] and Ct value remained linear. Primer-specific master mixes were prepared that contained SYBR Green, water, and the gene-specific primers and dispensed to a 96-well plate. Two microliters of the cDNA samples was then added to each well using an electronic repeater pipette (Rainin). The transcript levels of the reference gene, ACTIN2, did not vary by more than 1 Ct between treatments, suggesting that this is an appropriately stable transcript for normalization. All qRT-PCR values represent three biological replicates each containing three technical replicates. These biological replicates represent samples grown at three separate times, but with very carefully matched growth conditions, including medium, light, and time of day. The reference and target Ct numbers were entered into an Excel spreadsheet, where the efficiency-corrected relative expression, measured in fold increase over the untreated control, was calculated using an efficiency-corrected ΔΔCt formula (Pfaffl, 2001). The results presented in Figure 2 were confirmed by calculating fold change using an absolute quantification strategy similar to that described by Whelan et al. (2003; data not shown).

Root Tip Analysis of Flavonol Glycosides by Flow Injection Electrospray Mass Spectrometry

Roots were removed from the plates, and individual tips (approximately 5 mm in length) were excised, placed in Corning 2.0-mL self-standing microcentrifuge tubes, and immediately frozen and stored at −80°C. At the time of analysis, tubes were placed on ice and each root tip was suspended in 20 μL of extraction buffer (80% aqueous methanol, 1% acetic acid). The samples were then sonicated for 10 min in a sonicating water bath and then centrifuged (13,800_g_) for 2 min at room temperature. Ten microliters of the supernatant was then analyzed by flow injection using a TEMPO autosampler, in-line PEEK filter (0.5 μm), and nano multidimensional liquid chromatography unit (AB Sciex) interfaced with a 4000 QTrap linear ion-trap mass spectrometer (AB Sciex) via a MicroIon II nanospray source (AB Sciex).

Flow rate was maintained at 1.3 μL min−1 (80% aqueous methanol, 1% acetic acid). Precursor ion scans utilizing positive ion low-resolution mode were used to detect and quantitate molecules that, when fragmented within the mass spectrometer, yielded ions characteristic of kaempferol (m/z 287.1), quercetin (m/z 303.1), and isorhamnetin (m/z 317) derivatives. Parameters used for the precursor ion scans were 2.7 kV for the ion spray voltage, 16 V for the declustering potential, 11.5 V for the entrance potential, 2.5 V for the collision cell exit potential, interface heater set at 120°C, and curtain and nebulizer gases set to 10 and 27 (arbitrary units). Biological triplicates were analyzed in random order with blank runs between each sample. Scans corresponding to each precursor were collected for approximately 2.5 min and averaged. Peak areas corresponding to the most abundant flavonol glycosides (kaempferol-deoxyhexose-hexose, 595 m/z; quercetin-deoxyhexose-hexose, 611 m/z; isorhametin-rhamnose-hexose, 625 m/z) were then calculated using Analyst 1.4.2 (AB Sciex).

DPBA Staining and Quantification

Individual plants transferred to medium containing 1 μm IAA or 1 μm ACC for the indicated times were submerged in an aqueous solution containing 0.01% Triton X-100, 2.52 mg mL−1 DPBA, and placed on a rotary shaker at low speed for 7 min. The roots were then washed in deionized water for 7 min on the same shaker and mounted in deionized water between two coverslips. A Zeiss 710 LSCM (Carl Zeiss Microimaging) was used to excite the roots with 30% maximum laser power at 458 nm. DPBA fluoresces with distinct spectral properties in complex with kaempferol and quercetin (Neu, 1956; Sheahan and Rechnitz, 1992; Peer et al., 2001), with Q-DPBA having a longer wavelength fluorescence emission than K-DPBA. In Arabidopsis seedlings, DPBA fluorescence is largely due to binding to glycosylated flavonols, as aglycone flavonols are of low abundance or nondetectable in these tissues, as determined by HPLC analyses (Burbulis et al., 1996; Böttcher et al., 2008; Yonekura-Sakakibara et al., 2008). We optimized LSCM settings to spectrally separate the fluorescence of K-DPBA and Q-DPBA (which reflects the fluorescence of the glycosylated forms of kaempferol and quercetin) by capturing a window of the emission spectrum for each compound where they do not overlap. Fluorescence emission spectra were generated using the spectral imaging capabilities of the LSM 710 using the tt7-2 mutant and the tt4-2 mutant fed with kaempferol for K-specific fluorescence and quercetin for Q-specific fluorescence. In addition, isorhamnetin was added to the tt4 mutant, and an emission spectrum was captured. The constant dose of the flavonols added (5 μm) enables direct comparison of fluorescence quality and quantity (Fig. 3A), suggesting that kaempferol and quercetin are the major flavonols fluorescing when the wild type is stained with DPBA. The gain settings were selected to maximize the signal while eliminating cross-channel bleed-through from pure quercetin or K-DPBA in vitro and in vivo. These settings collect fluorescence at 475 to 504 nm for kaempferol and 577 to 619 nm for quercetin. Digital gain remained at 1 for kaempferol and was set at 0.3 for quercetin due to the stronger fluorescence of Q-DPBA relative to K-DPBA. This approach precludes absolute comparison of kaempferol and quercetin levels but can accurately measure relative increases in both. All micrographs within each figure (except for Fig. 4 as described below) were acquired using identical offset, gain, and pinhole settings using the same detectors.

Within each experiment and data presentation, gain was held constant. For images showing the flavonol accumulation patterns in Figure 4, A and C, gain was set at 650 for kaempferol and 515 for quercetin (Fig. 4A) to prevent saturation when DPBA staining increased in samples treated with IAA and ACC. For Figure 4C, the controls were imaged at higher gain, at 700 for kaempferol and 575 for quercetin, because the mutants had weaker fluorescence intensity. For the data in Table II, where both the wild type and mutants were imaged in the presence and absence of IAA and ACC, intermediate gain settings of 675 and 545 for kaempferol and quercetin, respectively, were used. To create a maximum projection of fluorescence from multiple focal planes, the 10× objective was used with a pinhole diameter of 2 airy units, and seven to 11 sections were collected with approximately 10% overlap. The stack was then rendered using the Zen software package (Carl Zeiss Microimaging) using a threshold value of 15%.

For quantification of DPBA fluorescence, the focal plane of maximum fluorescence intensity in the elongation zone was imaged. Postimaging, a box was drawn containing seven cells in two adjacent files at an equivalent distance from the apex. The average intensity of the fluorescence in this box was recorded. Fluorescence data were normalized and represented as fold change over untreated control. Therefore, the fluorescence is essentially reported with constant values per cell.

Construction of Transgenic Arabidopsis Plants Expressing CHSpro:CHS-GFP

The construct containing the CHS promoter, CHS coding region, and mGFP5 reporter was constructed using MultiSite Gateway Technology (Invitrogen). Amplification reactions were performed using PfuUltra polymerase (Stratagene) with primers (Integrated DNA Technologies) designed to introduce the appropriate TOPO or att sites (Supplemental Table S5). The AtCHS promoter region, defined as the 1,328 bp between the stop codon of the upstream gene and the start codon of AtCHS, was amplified from genomic wild-type (Columbia) DNA and inserted into the pDONR P4-P1R vector using the BP recombination reaction (Invitrogen). The CHS coding region was amplified from a construct previously generated in a modified pET32a(+) vector (Novagen; C. Dana and B.S.J. Winkel, unpublished data). Although these sequences were derived from the Landsberg erecta ecotype, the encoded amino acid sequence is identical to that of Columbia. The fragment was inserted into the pENTR/D-TOPO vector using the pENTR Directional TOPO Cloning Kit (Invitrogen). The coding sequences for mGFP5 were amplified from pAVA393 (von Arnim et al., 1998) and inserted into the pDONR P2R-P3 vector using the BP recombination reaction (Invitrogen). The integrity of each entry clone was confirmed by sequencing. The three entry vectors were recombined with the plant binary destination vector pB7m34GW (Flanders Institute for Biotechnology, Ghent University; Karimi et al., 2005) using the MultiSite Gateway LR reaction (Invitrogen).

The resulting multifragment construct was used to transform Agrobacterium tumefaciens strain GV3101 using a freeze-thaw method (Chen et al., 1994) and then introduced into tt4-11 (SALK 020583) plants using the floral dip method (Clough and Bent, 1998). Selection of transformants for resistance to Basta was carried out on T1 seedlings as described previously (Frazzon et al., 2007). Genomic DNA was extracted from putative transformants (Edwards et al., 1991) and analyzed by PCR using primers specific for the transgene.

Confocal Imaging of CHSpro:CHS-GFP and DR5:vYFP

Roots expressing CHSpro:CHS-GFP were transferred to 1 μm ACC or control medium, stained with 25 μg mL−1 propidium iodide, and excited with a 488-nm laser line attenuated to 20% power. GFP fluorescence was captured from 500 to 539 nm at a gain setting of 556. Propidium iodide fluorescence was captured from 589 to 719 nm at a gain setting of 395. A 10× objective was used, and consecutive z-slices were taken with the pinhole diameter set at 2 airy units and 10% overlap between sections. A maximum projection of these slices was computed using a threshold value of 15%. Quantification was carried out in the cortical cell layer of the elongation zone from seven cells from the same region of the root used for quantification of DPBA by averaging the intensity within this area.

DR5:vYFP was imaged in roots stained with 25 μg mL−1 propidium iodide by exciting the tissue with a 514-nm laser line attenuated to 50% power. YFP fluorescence was collected from 518 to 570 nm, and propidium iodide fluorescence was collected from 598 to 719 nm. The 10× objective was used with a pinhole diameter of 2 airy units. All images were captured with identical laser intensity, gain, and pinhole settings.

High-Resolution Gravitropism Analysis and Root Elongation Analysis

Five-day-old plants were transferred to control or ACC plates that had been acclimated to room temperature in a sterile hood to remove excess moisture and placed in a two-axis custom plate holder that was designed with two TSX-1D linear translation stages (Newport). The plate was positioned 13 cm from an Optem Macro video zoom lens (Qoptiq) attached to a Marlin firewire camera (Allied Vision Technologies). Roots were reoriented by turning the plate 90° under uniform light of 15 μmol m−2 s−1. Acquisition was automatically performed by the manufacturer’s software every 2 min at 80 pixels mm−1 resolution. Image stacks were compressed and transferred via ftp to a server at the University of Wisconsin for analysis.

We used a novel algorithm to analyze the tip angle during the gravitropic response. First, the root was isolated from the background of a grayscale image using the method of Otsu (1979), which finds a threshold value that accurately bins grayscale values into two categories: high intensity and low intensity. The result is a binary image with the root set to 1 and the background set to 0. The boundary of each binary root was traced, and the curvature was calculated along this path. Curvature along a path is defined as the ratio between the change in angle and the arc-length distance along the boundary. Gaussian convolution was used to smooth the derivatives used to calculate curvature to eliminate non-root-tip curvature maxima. The root tip was defined as the location where the curvature along the boundary was highest.

Once the coordinates of the root tip were defined, the binary image of the root was sampled within a disc with a radius of 50 pixels to calculate the tip angle. The (x, y) pairs that lie within the root were selected, and a principal component analysis was performed on the coordinates. The resulting first principal vector points in the direction tangent to the root tip. The angle between the first principal vector and the image horizon line is measured as the tip angle.

For elongation studies, plants were transferred to fresh control agar medium or medium containing 1 μm ACC with continued growth at 100 μmol m−2 s−1. At the indicated times after transfer, plates were scanned and were analyzed using ImageJ (National Institutes of Health) for root length.

Auxin Transport Assays

Basipetal auxin transport was assayed using 100 nm [3H]IAA applied in agar droplets as described (Lewis and Muday, 2009). After a 5-h incubation with the [3H]IAA-containing agar in contact with the root tip, the apical 2 mm was discarded and the radioactivity in the region extending from 2 to 7 mm basal to the site of application was determined using a Beckman LS6500 scintillation counter.

The following genes (with accession numbers) were examined in this study: CHS, AT5G13930; CHI, AT3G55120; F3H, AT3G51240; F3′H, AT5G07990; DFR, AT5G42800; FLS1, AT5G08640; EIN2, AT5G03280; ETR1, AT1G66340; TIR1, AT3G62980; MYB12, AT2G47460; PAP1, AT1G56650; and TTG1, AT5G24520.

Supplemental Data

The following materials are available in the online version of this article.

• Supplemental Figure S1. Absolute values of quercetin, kaempferol, and isorhamnetin in individual root tips after IAA and ACC treatment.
• Supplemental Figure S2. The expression of CHS, F3′H, FLS, and CHI is tissue layer and axial position specific.
• Supplemental Figure S3. IAA- and ACC-dependent increases in PAP1 and TTG1 transcription are lost in myb12.
• Supplemental Table S1. Relative expression of selected transcription factors in response to IAA and ACC treatment.
• Supplemental Table S2. Expression of flavonoid metabolism-linked transcription factors after treatment with IAA and ACC.
• Supplemental Table S3. Flavonoid pathway enzyme promoter regions contain auxin, ethylene, and MYB response elements.
• Supplemental Table S4. Sequences of primers used for qRT-PCR analysis of gene expression.
• Supplemental Table S5. Primers used for the generation of multifragment Gateway constructs.

Acknowledgments

We appreciate the assistance of Anita McCauley with the confocal imaging and her comments on the manuscript. We acknowledge the guidance of Rodrigo Velarde in optimizing the qRT-PCR experiments. We thank Tanya Falbel for introducing DR5:vYFP into the tt4 background. The Arabidopsis Biological Resource Center Stock Center (http://www.arabidopsis.org/abrc/) provided T-DNA insertion lines. Tools available on The Arabidopsis Information Resource (http://www.arabidopsis.org) and the Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/) were used for bioinformatic analysis of available microarray data.

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