The nuclear TIR1/AFB-Aux/IAA auxin pathway plays a crucial role in regulating plant growth and development. Specifically, the IAA17/AXR3 protein participates in Arabidopsis thaliana root development, response to auxin and gravitropism. However, the mechanism by which AXR3 regulates cell elongation is not fully understood.
We combined genetical and cell biological tools with transcriptomics and determination of auxin levels and employed live cell imaging and image analysis to address how the auxin response pathways influence the dynamics of root growth.
We revealed that manipulations of the TIR1/AFB-Aux/IAA pathway rapidly modulate root cell elongation. While inducible overexpression of the AXR3-1 transcriptional inhibitor accelerated growth, overexpression of the dominant activator form of ARF5/MONOPTEROS inhibited growth. In parallel, AXR3-1 expression caused loss of auxin sensitivity, leading to transcriptional reprogramming, phytohormone signaling imbalance and increased levels of auxin. Furthermore, we demonstrated that AXR3-1 specifically perturbs nuclear auxin signaling, while the rapid auxin response remains functional.
Our results shed light on the interplay between the nuclear and cytoplasmic auxin pathways in roots, revealing their partial independence but also the dominant role of the nuclear auxin pathway during the gravitropic response of Arabidopsis thaliana roots.

Introduction

Cell growth is a fundamental part of plant development and responses to external stimuli. The combination of cell division and cell elongation is the dominant factor in plant body formation. In the root tip, cells divide in the meristematic zone (Dolan et al., 1993) and then elongate in the elongation zone, driving the root growth (Beemster & Baskin, 1998). Many genetic and environmental factors determine root cell growth, including internal growth regulators – phytohormones (Vanstraelen & Benková, 2012).

One of the key hormones that affects root growth is auxin. Auxin is involved in many root developmental processes, from its role in meristem maintenance, through controlling the overall root architecture, to the formation of root hairs and lateral roots (reviewed in Overvoorde et al., 2010). Auxin gradient is associated with patterns of root cell proliferation, elongation, and differentiation. Its concentration is highest around the quiescent center, promoting cell stemness and division, and decreases gradually in the shootward direction, leading to cell differentiation (Grieneisen et al., 2007; Mähönen et al., 2014).

Apart from these processes, auxin exerts an inhibitory effect on root cell elongation, which was shown already in 1939 (Bonner & Koepfli, 1939). This effect is mediated by modifying cell wall extensibility, membrane depolarization and effects on intracellular signaling processes (Rayle & Cleland, 1992; reviewed in Majda & Robert, 2018). Furthermore, asymmetric redistribution of auxin in the gravistimulated root and its accumulation in the lower part of the root causes inhibition of cell elongation, resulting in bending of the root in the direction of gravity (Friml et al., 2002). How auxin governs such a wide range of responses occurring on long time scales (cell division and differentiation) and on short time scales (cell growth inhibition) is not completely understood. One explanation might be the involvement of the transcriptional and the rapid, nontranscriptional auxin signaling pathways (Dubey et al., 2023).

It is well established that auxin is perceived in the cell nucleus by the TIR1/AFB – Aux/IAA (Transport Inhibitor Response 1/Auxin F-Box - Auxin/Indole-3-Acetic Acid) co-receptor complex, representing the nuclear auxin pathway (Leyser, 2018). The TIR1/AFB proteins are part of an Skp-Cullin-F-box complex that drives degradation of the Aux/IAA proteins via the 26S proteasome. Upon high auxin levels, Aux/IAAs-mediated repressor recruitment is hindered, auxin response factors (ARFs) are released, resulting in modulation of transcription. This nuclear auxin pathway (NAP) plays a significant role in regulating plant growth and development (Leyser, 2018). In addition, it has been recently suggested that the adenylate cyclase activity of TIR1/AFBs is required for the function of NAP (Qi et al., 2022).

Auxin also triggers changes in plasma membrane potential, cytosolic Ca²⁺ spikes, and a rise in the apoplastic pH (reviewed in Dubey et al., 2021). According to the acid growth theory, an acidic pH enables cell wall extension through the regulation of the activity of cell wall-modifying enzymes (Rayle & Cleland, 1992). However, Serre et al. (2023) revealed that root surface pH does not fully correlate with local growth rates, showing distinct acidic and alkaline domains on the Arabidopsis root surface. They found the lowest pH values at the very tip of the root and in the maturation zone, with an alkaline domain in the transition zone and partially covering the early elongation zone. The establishment of these domains is modulated by components of the rapid auxin response (i.e. cytoplasmic auxin pathway – CAP), including the AFB1 auxin co-receptor, the Cyclic Nucleotide Gated Channel 14 (CNGC14), and the auxin influx carrier AUX1 (Auxin Resistant 1).

The change in apoplastic pH and the creation of pH domains are crucial during the gravitropic response (Monshausen et al., 2011; Shih et al., 2015; Serre et al., 2023). Upon gravistimulation, the alkaline domain on the lower nongrowing side of the root rapidly increased, while on the upper elongating side, the alkaline domain diminished, and the root surface became acidic.

In the root, a mutualistic interaction of auxin signaling pathways was shown. The synergistic functioning of the nuclear auxin pathway and the cytoplasmic AFB1 receptor for fast auxin responses (Prigge et al., 2020; Serre et al., 2021; Dubey et al., 2023) is essential for the regulation of root bending. However, their antagonistic effect is involved in the formation of lateral roots (Dubey et al., 2023). On the contrary, the transmembrane kinase 1-mediated apoplastic auxin pathway (AAP) acts antagonistically to the NAP in the regulation of root apoplastic pH or root growth inhibition (Li et al., 2021).

The semi-dominant axr3-1 Arabidopsis mutant harbors a stabilizing point mutation in the auxin binding domain II of the IAA17/AXR3 protein. This leads to severe growth and developmental defects, including agravitropic and auxin-insensitive roots (Leyser et al., 1996; Knox, 2003; Swarup et al., 2005; Lucas et al., 2008; Li et al., 2009; Mähönen et al., 2014) which fail to develop root hairs and lateral roots (Knox, 2003; Kim et al., 2006).

In this work, we exploit the mutant AXR3-1 protein as a tool to unravel how Aux/IAAs regulate root growth. By analyzing the line inducibly overexpressing AXR3-1, we show that the accumulation of stable AXR3-1 protein leads to a temporary root growth acceleration. To investigate the molecular network and physiological processes occurring in elongating root cells, we use pharmacological and genetical approaches and transcriptomic profiling. Our results provide insights into molecular mechanisms involved in regulation of cell elongation and expand our knowledge of relationships between the transcriptional and nontranscriptional auxin pathways in root.

Materials and Methods

Experimental material and generation of mutant plants

All lines used are in Arabidopsis thaliana (L.) Heynh. ecotype Columbia-0 (Col-0) background. We used the following lines: g1090::XVE>>AXR3-1-mCherry (iAXR3-1) (Mähönen et al., 2014), HS::AXR3-1 (Knox, 2003), iMPΔ (Gonzalez et al., 2021), DII-Venus (Brunoud et al., 2012), GPS2 sensor (Griffiths et al., 2023) and iaa17 (AT1G04250, SALK_065697C). iaa17 line was genotyped using the following primers: LP CGATTTTCCTCAAGTACGGTG and RP TTTCCTTCACTTGTGCTTTCG. g1090::XVE>>AXR3-1-mCherry were crossed with DII-Venus and GPS2 line. Lines g1090::XVE>>AXR3-1-ΔNLS-mScarlet, PIN2::XVE>>AXR3-1-ΔNLS-mScarlet, PIN2::GR-AXR3-1-mScarlet, PIN2::IAA17-mVenus and PIN2::AXR3-1-mScarlet were prepared in this study as described below.

Cloning strategy

GoldenBraid methodology (Sarrion-Perdigones et al., 2011) was used as a cloning strategy. For stable transgenic lines, we cloned coding sequence (CDS) of IAA17/AXR3 (AT1G04250) or AXR3-1 (88P → L substitution) driven by the PIN2 promoter (1.4 kb upstream of the AT5G57090) and fused with mVenus (PIN2::IAA17-mVenus) or mScarlet-I (Bindels et al., 2017, PIN2::AXR3-1-mScarlet) to C terminus. These constructs were terminated by 35S terminator and cloned into alpha1 vector.

For estradiol-inducible lines, a chimeric transcription activator XVE (Zuo et al., 2000) was cloned under the control of the PIN2 promoter (1.4 kb upstream of the AT5G57090) or g1090 promoter (Zuo et al., 2000) terminated by the RuBisCo terminator from Pisum sativum and cloned into alpha 1–1 vector. We cloned CDS of IAA17/AXR3, AXR3-1 or AXR3-1-ΔNLS (31K → E, 32R → S and 207R → G substitutions) downstream of the 4xLexA Operon driven by CaMV 35S minimal promoter (Sarrion-Perdigones et al., 2013) and terminated by the 35S terminator, fused with mScarlet-I (Bindels et al., 2017) to the C-terminal part and terminated by 35S terminator. For constructs containing glucocorticoid receptor (GR) (PIN2::GR-AXR3-1-mScarlet), GR was fused to N-terminus of AXR3-1. These constructs were cloned into the alpha1-3 vectors (Dusek et al., 2020). The alpha transcriptional units were then interspaced with matrix attachment regions (Dusek et al., 2020). Alpha vectors were combined with a Basta resistance cassette and cloned into the pDGB3omega1 binary vector (Sarrion-Perdigones et al., 2013).

All constructs were transformed into the Col-0 ecotype using the floral dip method (Clough & Bent, 1998). All used primers and sequences are listed in Supporting Information Table S1.

Growth conditions and treatments

Seeds were surface-sterilized with chlorine gas (Lindsey et al., 2017), and stratified for 2 d at 4°C. Seedlings were grown vertically on plates containing 1% (w/v) agar (Duchefa) with ½ Murashige and Skoog (MS, Duchefa, 0.5 g l⁻¹ MES, 1% (w/v) sucrose, pH 5.8 adjusted with 1 M KOH). Growth chamber conditions were 23°C by day (16 h), 18°C by night (8 h), light intensity of 120 μmol photons m⁻² s⁻¹, 60% humidity.

The chemicals used to prepare treatments are given in Table S2.

To treat the plants, 4–5 d old plants were transferred to treatment medium. Used concentrations and exact treatment times for specific experiments are given in the legend of each figure. In some cases, seeds were germinated on medium containing dexamethasone (DEX) or estradiol. Control medium always contained the respective mock treatment.

Microscopic imaging

For high-resolution imaging (including root growth rate measurement over time), vertical stage (von Wangenheim et al., 2017) Zeiss Axio Observer 7 coupled to a Yokogawa CSU-W1-T2 spinning disk unit with 50 μM pinholes and equipped with a VSHOM1000 excitation light homogenizer (Visitron Systems) was used. Images were acquired using the VisiView software (v.4.4.0.14; Visitron Systems, Puchheim, Germany). The roots were imaged with a Plan-Apochromat 10×/0.45 M27 and Plan-Apochromat 20×/0.8 M27 objectives. We used 515 nm laser for mVenus (excitation 515, emission 520–570 nm), 561 nm laser for mScarlet, mCherry and PI-stained samples (excitation 561, emission 582–636 nm) and 488 nm laser for DHE-/BES-H₂O₂-Ac-stained samples (excitation 488, emission 500–550 nm). For GPS2 sensor, sample were imaged by Zeiss LSM 880, C-Apochromat 40×/1.2 Imm Corr DIC objective. Fluorescence resonance energy transfer (FRET)/458 emission ratio was created as the ratio of FRET channel (excitation 548 nm, emission 525–579 nm) and the 458 channel (excitation 548, emission 463–517 nm) in a multichannel mode, in parallel, the yellow fluorescent protein channel was acquired (excitation 514, emission 525–579 nm).

Low-resolution imaging (gravitropic analysis, root growth rate measurement after 4 h) was performed by vertically placed flatbed scanner (Perfection V700; Epson, Suwa, Nagano, Japan) with the Epson Scanner software v3.9.2.1US.

Image analyses and measurements

Software ImageJ Fiji (Schindelin et al., 2012) was used for all image analysis.

For measuring root growth rate, distance between root tip positions in consecutive frames was calculated. Fiji plugin Correct 3D drift was used to stabilize the drift of the root tip.

To measure gravitropic bending, seedlings were transferred to plates containing the desired media. After 30 min for recovery (or 2 h for treatment), plates were turned 90° and imaged every 30 min. Analysis was done by Acorba software (Serre & Fendrych, 2022).

The size of the meristematic and elongation zone was measured on roots stained with 2.5 μM propidium iodide (PI) for 15 min. The elongation zone was measured on a time series, whereby the end of the elongation zone was set as the last growing cell. The size of the meristematic zone was measured as a distance from the root tip to the last isodiametric cell. The measurement of the meristematic zone in iMPΔ plants was more intricate, as adherence to this rule was not feasible. Instead, we used the end of the root cap as the boundary for the meristem.

The intensity of the fluorescent proteins was measured either as the intensity of the entire area from the root tip including the elongation zone in the case of mCherry or as the intensity of individual nuclei after removing the background intensity as in the case of the GPS2 sensor. To analyze the level of gibberellin, F1 cross GPS2 x g1090::XVE>>AXR3-1-mCherry were induced for 4 h. Gibberellin-dependent ratio change was calculated as FRET/405 fluorescence intensity.

Dihydroethidium (DHE, 5 mM stock in DMSO - dimethyl sulfoxide; Fisher Scientific, Waltham, MA, USA) and BES-H₂O₂-Ac (1 mM stock in DMSO; Wako, Richmond, VA, USA) were used to stain reactive oxygen species (ROS). After incubation in dark for 30 min, the signal intensity of epidermal cells was measured.

Microfluidic experiment for rapid growth inhibition was done as described in Serre et al. (2021).

Analyses of root pH profile was done as described in Serre et al. (2023).

Transcriptomic and gene expression analysis

To obtain transcriptomic data, RNA from 50 root tips (1–2 mm) of g1090::XVE>>AXR3-1-mCherry and HS::AXR3-1 were extracted following the protocol (Plant Total RNA Mini Kit; Qiagen). g1090::XVE>>AXR3-1-mCherry were transferred for 2 h on estradiol-containing medium and then harvested. HS::AXR3-1 plants were treated for 40 min with 37°C and harvested after another 80 min of recovery in cultivation room. Each of these lines had an appropriate control (treated in the same way as the mutant line). Col-0 was transferred to medium containing 10 nM IAA or 10 μM PEO-IAA or mock (ethanol) for 20 min. mRNA was prepared from total RNA, followed by Eukaryotic Strand-Specific Transcriptome Library preparation. RNA was sequenced by Illumina PE150 sequencing.

Quality of reads processed by the sequencing service provider was assessed by FastQC software. Transcript abundances (transcripts per million – TPM) were determined using Salmon v.1.3.0 (Patro et al., 2017) with parameters –validateMappings, –seqBias, –gcBias, –posBias, –numBootstraps 30. Reference index was built from Arabidopsis thaliana, TAIR10 cds library, v.20 101 214. Statistical evaluation and quality control of data analysis was done using Excel. Transcripts with the following parameters: SD TPM of all replicas/avg of all replicas ≤0.6 and log₂ fold change ≥ 1 (upregulated) or ≤ −1 (downregulated) were considered to be significantly differentially expressed. Gene ontology analysis was done using DAVID bioinformatics resources (Huang et al., 2009). Gene ontology (GO) with false discovery rate ≤ 0.05 and fold change ≥ 2 (upregulated) or ≤ −2 (downregulated) were considered to be significantly differentially expressed.

Selection of candidate genes and the phenotyping of mutants

Candidate genes for further analysis were selected based on the degree of change in their expression upon AXR3-1 overexpression and/or after IAA and PEO-IAA treatment. Genes showing root-specific expression were selected. Mutants were ordered from Nottingham Arabidopsis Stock Centre and subsequently verified by genotyping. The list of individual mutants including the primers used is in Table S3.

For the ploidy analysis, we followed a protocol from Urfus et al. (2021).

Liquid chromatography – mass spectrometry measurement

The endogenous phytohormones in root tips were determined according to Prerostova et al. (2021). In brief, c. 100 root tips of plants treated for 4 h with estradiol were collected into 100 μl acetonitrile/water (1/1, v/v), spiked with stable isotope-labeled internal standards (1 pmol per sample) and were homogenized with 1.5 mM zirconium beads using a FastPrep-24 homogenizer (MP Biomedicals, Irvine, CA, USA), and incubated at 4°C for 30 min. After centrifugation at 30 000 g for 20 min, the supernatant was concentrated to half of initial volume in vacuum concentrator and applied to SPE Oasis HLB 10 mg 96-well plate (Waters, Milford, MA, USA). The SPE 96-well plate was washed three times with 100 μl water, followed by elution with 100 μl 50% acetonitrile/water. Aliquot of 5 μl of the eluate was injected into the liquid chromatography-mass spectrometry (LC-MS) system consisting of UHPLC 1290 Infinity II (Agilent, Santa Clara, CA, USA) coupled to 6495 Triple Quadrupole mass spectrometer (Agilent). Mass spectrometry analysis was done in MRM mode, using isotope dilution method. Data acquisition and processing was done with Mass Hunter software B.08 (Agilent).

Statistical and graphic analyses

Each experiment was performed in a minimum of three biological replicates. To compare multiple samples, we used one-way ANOVA followed by Tukey HSD for normally distributed data. If data did not follow normal distribution, we used Kruskal–Wallis test followed by post hoc Dunn's test. To compare two sets of normally distributed samples, we used Student's t-test. P-value 0.05 > ns; *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001.

All data points in boxplots are shown as dots, the largest dot represents mean. Line graphs and data analysis was done in Microsoft Excel; boxplots were made by SuperPlotsOfData (Lord et al., 2020). Figures were assembled in Inkscape.

Results

Induction of AXR3-1 causes a promotion of root cell elongation

The severe phenotype of axr3-1 plants results from a long-term accumulation of the AXR3-1 protein, inevitably causing pleotropic secondary effects. To determine the primary effects of AXR3-1 protein accumulation in the Arabidopsis thaliana roots we used an estradiol-inducible g1090::XVE>>AXR3-1-mCherry line (iAXR3-1; Mähönen et al., 2014). Apart from the described disturbance of auxin sensitivity – root hair growth arrest, auxin insensitivity and agravitropic phenotype (Leyser et al., 1996; Knox, 2003; Mähönen et al., 2014) (Fig. 1a–c), AXR3-1 overexpression surprisingly triggered a temporary growth acceleration (Fig. 1d). Unlike the stable growth rate of the control line, iAXR3-1 plants displayed accelerated growth over several hours, reaching the maximal growth rate c. 4–6 h after the start of treatment. Later, the growth rate gradually decreased until root growth almost stopped (Fig. 1d). The effect of AXR3-1 induction could be detected 50 min after estradiol treatment, overcoming the auxin-induced growth inhibition (Fig. 1c). Interestingly, the growth acceleration preceded the detectable fluorescence of the AXR3-1-mCherry protein (Video S1), which is likely caused by the time required for the fluorescence protein to mature (Balleza et al., 2018).

Details are in the caption following the image — **Fig. 1**
Open in figure viewer PowerPoint

AXR3-1 overexpression results in altered root morphology and responses, including increased root cell elongation in *Arabidopsis thaliana*. Time series of Col-0 and *iAXR3-1* root after 0 h (white), 2 h (magenta), 4 h (green) and 6 h (yellow) of gravistimulation, 2 h of estradiol pretreatment (a); normalized gravitropic angle of *iAXR3-1* and Col-0, 2 h of estradiol pretreatment, normalized to the position of root tip at time point 0, n ≥ 11 (b); root growth rate of *iAXR3-1* and Col-0 after estradiol treatment and 10 nM IAA co-treatment, n = 10 (c); root growth rate of *iAXR3-1* and Col-0 after estradiol treatment, n ≥ 12 (d); *iAXR3-1* roots after 0, 4 and 10 h of estradiol treatment, arrows indicate the beginning and the end of the elongation zone (EZ) (e); size of the root meristematic zone (MZ), n ≥ 12 (f) and elongation zone, n ≥ 10 (g) of *iAXR3-1* plants after estradiol treatment; 10-d-old *iAXR3-1* plants grown on medium containing estradiol at different concentrations (h); error bars are SD in (b–d) and CI in (f, g); bars: (a, e) 50 μm; (h) 1 mm, asterisks indicate statistically significant differences based on one-way ANOVA followed by Tukey HSD; ***, P ≤ 0.001.

To test how the growth rate changes correlate with longitudinal zonation of the root tip, we determined the meristem and elongation zone size dynamics upon AXR3-1 induction. The meristem size decreased with the increasing time after induction (Fig. 1e,f). By contrast, the elongation zone size increased, reached the maximum size at 4 h and then gradually decreased (Fig. 1e,g), demonstrating that the root growth acceleration can be explained by the elongation zone size changes.

The growth acceleration occurred when iAXR3-1 plants were treated with at least 1 μM estradiol (Fig. S1a); however, iAXR3-1 roots were agravitropic already at 50 nm estradiol treatment (Fig. S1b). A concentration of 5 μM estradiol was used for all subsequent growth promotion experiments to achieve the highest effect on the mutant line with no apparent effect on the control line.

In addition to the short-term effects (h) of AXR3-1 induction, we analyzed its long-term effect on root growth and morphology. As expected, with the increasing concentrations of estradiol, the accumulation of AXR3-1-mCherry protein increased (Fig. S1c). This correlated with a decrease in root length, impaired formation of lateral roots (Fig. 1h) and root hairs (Fig. S1d). Already the low estradiol concentration treatment resulted in shorter, agravitropic roots that lacked lateral roots (Figs 1h, S1b) indicating different sensitivity of various auxin-related processes to AXR3-1 levels.

When we analyzed the iaa17/axr3 loss of function mutant, we did not detect differences in root growth rate, response to gravistimulation or auxin sensitivity when compared to the Col-0 control line (Fig. S1e–g). As IAA17/AXR3 is one of 29 Aux/IAA proteins (Reed, 2001), this results likely from the redundancy within this protein family.

Taken together, these results suggest the role of AXR3-1 in regulation of the dynamics of cell expansion and longitudinal zonation in the Arabidopsis root.

Growth acceleration and auxin insensitivity require nuclear localization of AXR3-1

The AFB1 auxin co-receptor and several Aux/IAA proteins localize not only to the nucleus, but also partially to the cytoplasm (Zhang et al., 2019; Prigge et al., 2020), and the AFB1-dependent rapid signaling that controls root cell elongation occurs in the cytoplasm (Prigge et al., 2020; Dubey et al., 2023). We, therefore, tested whether AXR3-1 functions outside the nucleus and participates in the cytoplasmic rapid auxin response. First, we expressed the wild-type IAA17/AXR3 and its stabilized form AXR3-1 under the control of the strong ubiquitous g1090 promoter (Ishige et al., 1999) and the PIN2 promoter, specific for the lateral root cap, epidermis and cortex root cells. Both protein forms localized exclusively to the nucleus (Fig. 2a), in agreement with Ouellet et al. (2001), and corresponding to the presence of the bipartite nuclear localization signal (NLS) in IAA17/AXR3 sequence (Abel et al., 1994). To reveal a potential role of a minor cytoplasmic fraction of IAA17/AXR3, we mutated the bipartite NLS of AXR3-1 using the 31K → E, 32R → S and 207R → G substitutions, creating the AXR3-1ΔNLS (Fig. 2b; Ouellet et al. (2001)). AXR3-1ΔNLS protein localized to the cytoplasm (Fig. 2c). In contrast to AXR3-1, the expression of AXR3-1ΔNLS in two independent homozygote lines did not interfere with auxin sensitivity (Figs 2d, S2a) or gravitropic response (Fig. 2g), and it did not affect the root phenotype in long-term treatment (Fig. S2b). Moreover, the expression of AXR3-1ΔNLS did not lead to growth acceleration. Instead, AXR3-1ΔNLS exerted a mild negative effect on root growth (Figs 2d, S2a).

In addition, we fused the AXR3-1 protein to the glucocorticoid receptor fragment (GR; Schmitt & Stunnenberg, 1993) to allow for a DEX-dependent relocalization from cytoplasm to the nucleus. In mock conditions (DMSO), the signal was present in the cytoplasm (Fig. 2e). Compared to the control, root growth was mildly negatively affected by AXR3-1 localized in cytoplasm (Fig. 2f). However, after the addition of DEX, the protein moved to the nucleus (Fig. 2e) and triggered a growth stimulation, auxin insensitivity (Fig. 2f) and an agravitropic phenotype (Fig. 2h). In agreement, plants grown on DEX-containing medium for 5 d were agravitropic (Fig. S2c). Notably, the expression of the positive control – the AXR3-1 protein localized in the PIN2 domain – led to growth acceleration and auxin insensitivity as expected. However, the effect on root growth was weaker than in the original iAXR3-1 plants, where the expression is driven by the g1090 promoter.

These results clearly show that growth stimulation caused by AXR3-1 overproduction requires AXR3-1 protein localized in the nucleus. This indicates that the observed growth phenotype depends on AXR3-1-mediated transcriptomic changes, rather than its involvement in cytoplasmic auxin response.

The growth promotion in iAXR3-1 correlates with transcriptional reprogramming

To identify the genes responsible for the growth stimulation after induction of AXR3-1, we analyzed transcriptomic profiles of root tips of two AXR3-1 inducible lines: estradiol-inducible iAXR3-1 and the heat shock-inducible HS::AXR3-1 (Knox, 2003). In addition, to identify genes regulated by auxin signaling, we obtained transcriptome of Col-0 root tips treated for 20 min with indole-3-acetic acid (IAA) and auxin signaling inhibitor (PEO-IAA, Nishimura et al., 2009).

In the resulting dataset, we could detect the IAA17/AXR3 gene as one of the most upregulated genes in iAXR3-1 roots; expression of heat shock proteins was upregulated after heat shock as expected; and the known auxin-inducible genes were induced in the IAA-treated roots (Fig. S3a). The heat shock induction, however, did not function efficiently in one of the replicas; therefore, we omitted the HS::AXR3-1 results from further analysis.

Pairwise comparison with the Col-0 control identified 906 differentially expressed genes in iAXR3-1, 160 of them were also differentially expressed in the IAA treatment (Dataset S1). We focused on genes downregulated in iAXR3-1 and with altered expression after IAA treatment. Further, we narrowed the candidate gene list based on their expression and/or described role in the root (using TAIR – Berardini et al., 2015). We obtained a set of 16 candidate genes potentially responsible for the iAXR3-1 phenotype. We isolated the respective mutants and analyzed their root growth phenotypes and additional auxin-related processes. We discovered several auxin-dependent phenotypic deviations: mutant in MIZU KUSSEI-LIKE showed a shorter root (Fig. S3b), the mutant in PECTIN METHYLESTERASE 41 (PME41) showed higher root hair density (Fig. S3c) and mutant in AUXIN UPREGULATED F-BOX PROTEIN1 (AUF1) displayed a higher number of lateral roots (Fig. S3d). The summary of these experiments and results can be found in Table S4. None of the tested mutants, however, showed any defects in auxin-regulated root growth.

Further, the dataset pointed out a significant downregulation of siamese related1 and siamese, known cyclin-dependent kinase inhibitors, that regulate endoreduplication in plants (Fig. S3e). Therefore, we compared the ploidy state of root tips of iAXR3-1 and Col-0 2 h and 20 h after estradiol treatment. The results, however, revealed no obvious difference in ploidy between the control and the iAXR3-1 line (Fig. S3f).

The fact that the screening of the selected mutants did not reveal a single gene that would control the rapid growth promotion suggests that the regulation is more complex, and it involves groups of genes or processes. Therefore, we performed the GO analyses (DAVID – Huang et al., 2009). We divided the differentially expressed genes in iAXR3-1 roots into functional groups based on their molecular and biological function, cell compartment, or the pathway in which they are included (Fig. 3a).

One of the intriguing observations is the significant overrepresentation of genes linked with ROS (Fig. 3a), particularly peroxidases. Reactive oxygen species have been shown to be involved in defining the border between meristematic and elongation zone (Tsukagoshi et al., 2010) and regulate cell wall extensibility (reviewed in Kärkönen & Kuchitsu, 2015). We, therefore, analyzed the levels of superoxide and hydrogen peroxide in induced iAXR3-1 roots using the DHE and BES-H₂O₂-Ac staining, respectively. We observed increased levels of both analyzed types of ROS in the rapidly elongating iAXR3-1 roots in comparison to controls (Fig. 3b). We could, however, not fully prevent the effect of iAXR3-1 on root elongation by increasing or decreasing ROS levels by hydrogen peroxide (H₂O₂) or the ROS scavenger N-acethyl-1-cysteine, respectively (Fig. S3g,h), neither could we stimulate cell elongation by applying increasing concentrations of H₂O₂ to Col-0 roots (Fig. S3i).

Together, these data suggest that modulated ROS level on its own cannot explain the increased cell elongation in iAXR3-1.

AXR3-1 triggers auxin overproduction

Notably, GO terms related to phytohormones were significantly enriched, including genes related to brassinosteroids, gibberellins, ethylene, and auxin (Fig. 3a; Dataset S1). Application of bioactive brassinolide or brassinosteroid synthesis inhibitor brassinazole resulted in the same growth response in iAXR3-1 and the control line (Fig. S4a). However, root elongation of the iAXR3-1 line was less sensitive to ethylene (Fig. S4b) and its precursor (Fig. S4c), confirming the interconnection of the auxin and ethylene signaling pathways (Růžička et al., 2007).

To address the role of gibberellin in the growth stimulation of iAXR3-1 roots, we biochemically altered gibberellin level in the root. The application of gibberellic acid 3 (GA3) and gibberellic acid 4 (GA4) without previous inhibition of gibberellin biosynthesis had no effect on root growth, regardless of the genotype we studied (Fig. S4d), corresponding to the published literature (Tanimoto, 1987; Rieu et al., 2008; Li et al., 2015). Inhibition of gibberellin biosynthesis using two different inhibitors (paclobutrazol – PAZ and uniconazole – Uni, Izumi et al., 1988) led to a decrease in root growth rate (Figs 3c, S4e) in both control and iAXR3-1 lines, already after 4 h of treatment. Interestingly, exogenously added GA3/GA4 rescued the growth rate of PAZ/Uni treated iAXR3-1 roots. By contrast, GA could not completely recover the growth rate after 4 h in PAZ/Uni treated Col-0 roots (Figs 3c, S4e). From this data, we conclude that iAXR3-1 plants are hypersensitive to gibberellin. Intriguingly, the expression of gibberellin catabolic and gibberellin synthetic genes was downregulated in iAXR3-1 (Fig. S4f), suggesting altered levels of gibberellin in iAXR3-1 roots. We, therefore, introduced the GPS2 – GIBBERELLIN PERCEPTION SENSOR 2 line into the iAXR3-1 to estimate gibberellin levels (Griffiths et al., 2023). We could, however, not detect any difference between gibberellin levels in iAXR3-1 and control roots (Fig. 3d). Nevertheless, it should be noted that GPS2 sensor's partial reversibility (Rizza et al., 2017; Griffiths et al., 2023) makes it challenging to detect a decrease in gibberellin levels during our short monitoring period.

The most overrepresented GO categories in iAXR3-1 roots were related to auxin, and in addition to auxin signaling genes, genes involved in auxin biosynthesis were upregulated after AXR3-1 induction (Fig. 3a). To approximate auxin levels, we crossed iAXR3-1 plants with the negative auxin reporter DII-Venus line (Brunoud et al., 2012). Upon expression of AXR3-1 protein, the DII-Venus signal rapidly disappeared (Fig. 3e), indicating increased auxin level in the AXR3-1-expressing cells. In accordance with these results, LC-MS measurement of the hormones level in the root tips confirmed a significant increase in the level of IAA and its metabolites in iAXR3-1 roots (Fig. 3f).

The results provide evidence that induction of AXR3-1 protein in the root led to increased auxin biosynthesis and its accumulation in root cells. In the iAXR3-1, the perturbed auxin response disturbs auxin homeostasis that probably further leads to imbalances in other phytohormonal pathways and might further influence root growth mediated by the other auxin perception pathways.

Transcriptional activator ARF5/MP inhibits root cell elongation

The fact that we were unable to attribute the induction of cell elongation in iAXR3-1 to an individual gene or a cellular process hints at the explanation that the cell elongation process is controlled in parallel by multiple genes the expression of which is altered in the induced line. The AXR3 protein is a negative regulator of auxin-induced transcription mediated by the activator ARF proteins (Gray et al., 2001; Pierre-Jerome et al., 2016), the logical explanation is that the activator ARFs and their downstream targets negatively regulate cell elongation in the root elongation zone. To test this hypothesis, we took advantage of the irrepressible version of ARF5/MONOPTEROS, lacking domain enabling Aux/IAAs binding (ARF5Δ/MPΔ) (Gonzalez et al., 2021). We used estradiol-inducible iMPΔ and monitored the dynamics of root cell elongation and longitudinal developmental zonation. The inducible expression of a MPΔ triggered a progressive inhibition of root growth that was detectable c. 90 min after induction, and that caused an almost complete root growth inhibition 4 h after the treatment (Fig. 4a,b). This is an almost perfectly opposite effect compared to the iAXR3-1 induction. Similarly to iAXR3-1 line, roots of iMPΔ lost the gravitropic response (Fig. 4a), however, as the roots almost stopped growing, gravitropism was difficult to score. Growth inhibition in iMPΔ line was caused by a rapid and substantial reduction in the size of the elongation zone (Fig. 4c,d). Additionally, the size of iMPΔ meristematic zone size did not decrease (Fig. 4e).

These results demonstrate that auxin via the activator ARFs and their downstream target genes negatively regulate the elongation of root cells. This pathway is perturbed in the iAXR3-1, leading to excessive cell elongation and perturbation of the longitudinal root zonation.

Rapid auxin responses are retained in iAXR3-1 mutant

As we showed, NAP inhibits root cell elongation through transcription of genes by the activator ARF5/MP and its inhibitor IAA17/AXR3. At the same time, CAP regulates cell expansion and alkalinization of the apoplast by the AFB1-CNGC14 module (Dubey et al., 2023; Serre et al., 2023). Do these pathways depend on each other? Is the rapid module still active in the induced iAXR3-1 plants?

Interestingly, chemical inhibition of auxin biosynthesis by the auxin biosynthesis inhibitor L-kynurenine (L-kyn, He et al., 2011) led to increased root growth rate of control plants and it triggered further growth stimulation also in iAXR3-1 line (Fig. 5a), demonstrating that iAXR3-1 roots maintained the ability to partially respond to endogenous auxin. To test whether the expression of AXR3-1 inhibits the rapid nontranscriptional root response to IAA, we used a microfluidic system allowing the change of control medium to treatment medium in a few seconds (Serre et al., 2021). Similarly to the control, iAXR3-1 roots responded to addition of auxin by a rapid reduction of growth rate, even though the resulting growth rate was still higher than in the control (Fig. 5b). Using an alternative method of growth rate analysis after a 20 min IAA treatment, we confirmed that iAXR3-1 roots retain the rapid auxin response (Fig. S4g).

Next, we focused on analyzing the longitudinal root surface pH profile. To study the effect of altered NAP on root pH profile, we visualized root surface pH of vertically growing iAXR3-1 and iMPΔ 4 h after induction using a pH-sensitive fluorescent dye fluorescein-5-(and-6)-sulfonic acid (Serre et al., 2023). Overexpression of AXR3-1 led to root surface acidification, whereas iMPΔ roots showed a more alkaline surface pH. These results fit the increased and decreased elongation rates in the iAXR3-1 and iMPΔ roots, respectively. Intriguingly, in both cases, the formation of an alkaline domain in the root transition zone, that is the hallmark of the activity of the AFB1-dependent CAP, was preserved (Fig. 5c,d), albeit the alkaline domain was less apparent in the iAXR3-1 roots. Since the formation of a pH gradient on the two sides of the root occurs during the gravitropic response, we measured the pH profile of iAXR3-1 roots after gravistimulation. In contrast to control, gravistimulated iAXR3-1 failed to create a pH gradient between the upper and lower root side, which is consistent with their agravitropic growth (Fig. 5e,f).

Our data showed that in vertically growing plants, the rapid response pathway (i.e. CAP) can still function independently on the NAP. However, the transcriptional auxin response pathway dominates the overall elongation of the root, and its function is strictly required for the gravitropic response of the roots.

Discussion

Auxin homeostasis and its perception is essential for optimal plant growth and development. The pool of auxin is maintained by various mechanisms, including regulation of auxin biosynthesis, conjugation, and transport (reviewed in Ljung et al., 2002). Accumulation of AXR3-1 protein interferes with auxin perception and ARF-modulated gene expression, including the negative transcriptional feedback mechanism controlling the auxin homeostasis (Suzuki et al., 2015). Our results suggest that root cells interpret the AXR3-1 accumulation as low auxin levels, and to re-establish the proper auxin level, increase the production of auxin. In parallel, the loss of transcriptional response to auxin triggers excessive cell elongation (Fig. 6). Moreover, as auxin gradient regulates root cell division, expansion, and differentiation (Sabatini et al., 1999; Mähönen et al., 2014), disruption of auxin transcriptional response in iAXR3-1 roots results in disbalance between cell division and elongation, leading eventually to growth arrest. These results indicate that endogenous auxin maintains optimal root growth rate and that the potential of root cells to elongate is not fulfilled under normal physiological conditions.

Reduction in the overall growth rate of axr3-1 mutants (Leyser et al., 1996; Knox, 2003; Mähönen et al., 2014) is consistent with the observed long-term effect of AXR3-1 expression. Hitherto, the growth promotion effect of short-term AXR3-1 induction has not been reported (Mähönen et al., 2014). To investigate the possible growth promotion effect of other stabilized Aux/IAAs (Knox, 2003), it would be necessary to focus on a shorter duration of their expression. Additionally, the observed growth arrest phenotype corresponds to findings in published studies, such as those involving auxin biosynthesis mutant (Zhao et al., 2001) or mutants with stable expressions of other AUX/IAA genes (Arase et al., 2012). The differences in sensitivity of individual processes can be explained by the wide range of auxin action, including spatial and temporal aspects and different sensitivity of individual cells to auxin (Sauer et al., 2013; Caumon & Vernoux, 2023). Furthermore, according to our results, the effect on both root cell elongation and gravitropism is more significant if AXR3-1 is expressed in all root cell types, indicating the contribution of inner cell types to growth control by auxin.

In contrast to the findings obtained using transient expression system (Zhang et al., 2019), we observed solely nuclear localization of AXR3-1. As shown before for the AXR3-1 effect on development (Ouellet et al., 2001), the root growth acceleration and disruption of gravitropism require a nuclear-localized AXR3-1 protein. Interestingly, we observed a weak but consistent effect of cytoplasm-localized AXR3-1 on growth rate. It is intriguing to speculate that the accumulation of the protein in the cytoplasm interferes with the AFB1-dependent auxin response pathway (Prigge et al., 2020; Dubey et al., 2023).

Increased sensitivity of iAXR3-1 to GA treatment, the crucial role of gibberellins in regulating root cell expansion (Fu & Harberd, 2003; Ubeda-Tomás et al., 2008; Tanimoto, 2012), and their accumulation in the elongation zone of the root (Shani et al., 2013; Rizza et al., 2017, this study) makes them a compelling candidate for co-regulators of AXR3-1 growth stimulation. Since inhibition of GA biosynthesis from the initial stages of AXR3-1 induction did not lead to complete prevention of growth stimulation, our results favor the explanation that the altered sensitivity to GA is a secondary effect of the loss of sensitivity to auxin and disruptions in NAP. The ability to perceive auxin and the consequent capability to regulate its homeostasis are essential for maintaining proper crosstalk with other hormones, as demonstrated in our case by a documented alteration in sensitivity to gibberellins.

Reactive oxygen species seemed to be one of the plausible candidates for the regulation of AXR3-1-dependent growth acceleration. Our results did not prove that hydrogen peroxide application on its own could accelerate root growth. However, it is questionable to what extent pharmacological treatment corresponds to ROS accumulating in different cellular compartments at various concentrations required for the creation of specific signals under physiological conditions (Mhamdi & Van Breusegem, 2018). Although we demonstrated an increased level of ROS in iAXR3-1 roots, this might be associated with a stress response caused by excessive cell elongation (Sharma et al., 2012). Nevertheless, ROS are involved in modulation of cell wall properties (reviewed in Kärkönen & Kuchitsu, 2015). It is a question of future research to investigate AXR3-1-dependent modulation of the cell wall and the possible involvement of ROS.

Since plants overproducing AXR3-1 are still able to perceive chemical reduction of endogenous auxin level and retain nontranscriptional rapid response to auxin, it is obvious that their capability to perceive auxin is not completely diminished.

Additionally, auxin-modulated apoplastic pH changes serve as one of the drivers of growth (Monshausen et al., 2011; Barbez et al., 2017; Serre et al., 2023), by the regulation the activity of cell wall proteins (Rayle & Cleland, 1992). Our results show the tight correlation of root surface pH with the overall root growth rate: while alkaline pH corresponds to slowed growth of iMPΔ, surface acidification correlates with growth promotion of iAXR3-1. The mechanism of how auxin alkalinizes the apoplast in roots remains unresolved, both for the AFB1-dependent rapid response pathway (i.e. CAP) and the ARF-dependent transcriptional pathway. Our results are consistent with the previously published results (Li et al., 2021; Serre et al., 2023), but hint to that the mechanism of root surface pH control is dramatically different in roots and aboveground parts, where auxin via the TIR1/AFB-Aux/IAA pathway activates AHA proton pumps (Haruta et al., 2015; Fendrych et al., 2016).

What is the molecular link between NAP and CAP, and what is its biological significance in the root? The preservation of the rapid auxin response and the alkaline surface pH domain in the transition zone of iAXR3-1 roots is consistent with the model that CAP controls surface pH and the rapid auxin growth responses independently on NAP (Serre et al., 2021, 2023; Dubey et al., 2023). On the other hand, the ARF-dependent transcriptional responses clearly dominate the overall root surface pH. Based on the retained nontranscriptional pathway in iAXR3-1 plants, one would expect the pH profile to be maintained during gravistimulation, which is not the case. This might indicate that the rapid nontranscriptional auxin response during gravistimulation depends on the functional NAP. A more plausible explanation might be the following: we demonstrated that iAXR3-1 overproduces auxin in all tissues. Therefore, the essential requirement of the Cholodny-Went model – the asymmetric auxin distribution – is likely not fulfilled in the iAXR3-1 roots.

In summary, our data highlights the crucial role of the NAP components – ARF5/MP and IAA17/AXR3 in regulating Arabidopsis root growth. Sudden overexpression of AXR3-1 leads to losing the sensitivity of NAP to auxin, resulting in transcriptional reprogramming, followed by secondary effects such as changes in the root's balance in phytohormones and accumulation of ROS. These processes collectively result in uncontrolled, accelerated root cell elongation in iAXR3-1 (Fig. 6). This disruption in the balance between elongation and proliferation exhausts the meristem and halts root growth. Additionally, intact CAP, with the contribution of NAP and antagonistic effect of AAP (Li et al., 2021) modulate apoplastic pH.

Our findings offer insights into the connections among distinct auxin signaling pathways, their interaction in the regulation of root cell elongation and underscore the significance of examining their impacts over varying timeframes.

Acknowledgements

MF team received support from the European Research Council (Grant no. 803048). MK was supported by Charles University Grant Agency (Grant no. 337021). KM acknowledges the support of the Czech Science foundation (GACR 23-07813S). AMJ and AR were supported by the Gatsby Charitable trust (GAT3395) and European Research Council under the European Union's Horizon 2020 research and innovation program (Grant no. 759282). The authors are grateful to Jan Petrášek, Matouš Glanc for critical reading the manuscript and for discussion and Eva Medvecká for lab support, Bára Kolrosová for help with the collection of material for the transcriptomic experiment and professor Bargmann for kindly providing us with iMPΔ seeds. Confocal laser scanning microscopy performed in the Microscopy Core Facility of Faculty of Science was co-financed by the Czech-BioImaging large RI project LM2023050.

Competing interest

None declared.

Author contributions

MF and MK conceived and designed the experiments and wrote the manuscript. MK performed the experiments. KM analyzed transcriptomic data. PID performed LC-MS assay. AR and AMJ provided GPS2 sensor.

Open Research

Data availability

Rough sequencing data and quantification tables are available through GEO Series accession no. GSE227820 at the Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/). Source data were deposited at Zenodo, doi: 10.5281/zenodo.10531838.

Supporting Information

Filename

Description

nph19557-sup-0001-DatasetS1.xlsxExcel 2007 spreadsheet , 26.6 MB

Dataset S1 Raw transcriptomic data of iAXR3-1 and plants treated with auxin.

nph19557-sup-0002-SupInfo.pdfPDF document, 1 MB

Fig. S1 The AXR3-1 phenotype exhibits dose dependency.

Fig. S2 AXR3-1 functions within the nucleus.

Fig. S3 Potential candidate genes or processes responsible for iAXR3-1 phenotype.

Fig. S4 Impact of changes in various hormone levels on iAXR3-1 phenotype.

Table S1 List of primers and sequences used for cloning.

Table S2 List of chemicals used for treatments.

Table S3 List of mutants and primers used for experiments given in Table S4.

Table S4 List of analyzed candidates including their enrichment in iAXR3-1 or in IAA-treated Col-0 roots and the list of experiments.

nph19557-sup-0003-VideoS1.aviAVI video, 18.3 MB

Video S1 Root with inducible expression of AXR3-1 after estradiol treatment.

Please note: Wiley is not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.

References

Abel S, Oeller PW, Theologis A. 1994. Early auxin-induced genes encode short-lived nuclear proteins. Proceedings of the National Academy of Sciences, USA 91: 326–330.
10.1073/pnas.91.1.326
CASPubMedWeb of Science®Google Scholar
Arase F, Nishitani H, Egusa M, Nishimoto N, Sakurai S, Sakamoto N, Kaminaka H. 2012. IAA8 involved in lateral root formation interacts with the TIR1 auxin receptor and ARF transcription factors in Arabidopsis. PLoS ONE 7: e43414.
10.1371/journal.pone.0043414
CASPubMedWeb of Science®Google Scholar
Balleza E, Mark Kim J, Cluzel P. 2018. Systematic characterization of maturation time of fluorescent proteins in living cells. Nature Methods 15: 47–51.
10.1038/nmeth.4509
CASPubMedWeb of Science®Google Scholar
Barbez E, Dünser K, Gaidora A, Lendl T, Busch W. 2017. Auxin steers root cell expansion via Apoplastic PH regulation in Arabidopsis thaliana. Proceedings of the National Academy of Sciences, USA 114: E4884–E4893.
10.1073/pnas.1613499114
CASPubMedWeb of Science®Google Scholar
Beemster GTS, Baskin TI. 1998. Analysis of cell division and elongation underlying the developmental acceleration of root growth in Arabidopsis thaliana. Plant Physiology 116: 1515–1526.
10.1104/pp.116.4.1515
CASPubMedWeb of Science®Google Scholar
Berardini TZ, Reiser L, Li D, Mezheritsky Y, Muller R, Strait E, Huala E. 2015. The Arabidopsis information resource: making and mining the ‘gold standard’ annotated reference plant genome. Genesis 53: 474–485.
10.1002/dvg.22877
CASPubMedWeb of Science®Google Scholar
Bindels DS, Haarbosch L, van Weeren L, Postma M, Wiese KE, Mastop M, Aumonier S, Gotthard G, Royant A, Hink MA et al. 2017. MScarlet: a bright monomeric red fluorescent protein for cellular imaging. Nature Methods 14: 53–56.
10.1038/nmeth.4074
CASPubMedWeb of Science®Google Scholar
Bonner J, Koepfli JB. 1939. The inhibition of root growth by auxins. American Journal of Botany 26: 557–566.
10.1002/j.1537-2197.1939.tb09319.x
CASWeb of Science®Google Scholar
Brunoud G, Wells DM, Oliva M, Larrieu A, Mirabet V, Burrow AH, Beeckman T, Kepinski S, Traas J, Bennett MJ et al. 2012. A novel sensor to map auxin response and distribution at high spatio-temporal resolution. Nature 482: 103–106.
10.1038/nature10791
CASPubMedWeb of Science®Google Scholar
Caumon H, Vernoux T. 2023. A matter of time: auxin signaling dynamics and the regulation of auxin responses during plant development. Journal of Experimental Botany 74: 3887–3902.
10.1093/jxb/erad132
CASPubMedWeb of Science®Google Scholar
Clough SJ, Bent AF. 1998. Floral dip: a simplified method for agrobacterium-mediated transformation of Arabidopsis thaliana. The Plant Journal 16: 735–743.
10.1046/j.1365-313x.1998.00343.x
CASPubMedWeb of Science®Google Scholar
Dolan L, Janmaat K, Willemsen V, Linstead P, Poethig S, Roberts K, Scheres B. 1993. Cellular organisation of the Arabidopsis thaliana root. Development 119: 71–84.
10.1242/dev.119.1.71
CASPubMedWeb of Science®Google Scholar
Dubey SM, Han S, Stutzman N, Prigge MJ, Medvecká E, Platre MP, Busch W, Fendrych M, Estelle M. 2023. The AFB1 auxin receptor controls the cytoplasmic auxin response pathway in Arabidopsis thaliana. Molecular Plant 16: 1120–1130.
10.1016/j.molp.2023.06.008
CASPubMedWeb of Science®Google Scholar
Dubey SM, Serre NBC, Oulehlová D, Vittal P, Fendrych M. 2021. No time for transcription—rapid auxin responses in plants. Cold Spring Harbor Perspectives in Biology 13: a039891.
10.1101/cshperspect.a039891
CASPubMedWeb of Science®Google Scholar
Dusek J, Plchova H, Cerovska N, Poborilova Z, Navratil O, Kratochvilova K, Gunter C, Jacobs R, Hitzeroth II, Rybicki EP et al. 2020. Extended set of goldenbraid compatible vectors for fast assembly of multigenic constructs and their use to create geminiviral expression vectors. Frontiers in Plant Science 11: 1–11.
10.3389/fpls.2020.522059
PubMedWeb of Science®Google Scholar
Fendrych M, Leung J, Friml J. 2016. TIR1/AFB-Aux/IAA auxin perception mediates rapid cell wall acidification and growth of Arabidopsis hypocotyls. eLife 5: 1–19.
10.7554/eLife.19048
Web of Science®Google Scholar
Friml J, Wiśniewska J, Benková E, Mendgen K, Palme K. 2002. Lateral relocation of auxin efflux regulator PIN3 mediates tropism in Arabidopsis. Nature 415: 806–809.
10.1038/415806a
PubMedWeb of Science®Google Scholar
Fu X, Harberd NP. 2003. Auxin promotes Arabidopsis root growth by modulating gibberellin response. Nature 421: 740–743.
10.1038/nature01387
CASPubMedWeb of Science®Google Scholar
Gonzalez JH, Taylor JS, Reed KM, Clay Wright R, Bargmann BOR. 2021. Temporal control of morphogenic factor expression determines efficacy in enhancing regeneration. Plants 10: 2271.
10.3390/plants10112271
CASGoogle Scholar
Gray WM, Kepinski S, Rouse D, Leyser O, Estelle M. 2001. Auxin regulates SCFTIR1-dependent degradation of AUX/IAA proteins. Nature 414: 271–276.
10.1038/35104500
CASPubMedWeb of Science®Google Scholar
Grieneisen VA, Jian X, Marée AFM, Hogeweg P, Scheres B. 2007. Auxin transport is sufficient to generate a maximum and gradient guiding root growth. Nature 449: 1008–1013.
10.1038/nature06215
CASPubMedWeb of Science®Google Scholar
Griffiths J, Rizza A, Tang B, Frommer WB, Jones AM. 2023. Gibberellin perception sensors 1 and 2 reveal cellular GA dynamics articulated by COP1 and GA20ox1 that are necessary but not sufficient to pattern hypocotyl cell elongation. BioRxiv. doi: 10.1101/2023.11.06.565859.
Google Scholar
Haruta M, Gray WM, Sussman MR. 2015. Regulation of the plasma membrane proton pump (H⁺–ATPase) by phosphorylation. Current Opinion in Plant Biology 28: 68–75.
10.1016/j.pbi.2015.09.005
CASPubMedWeb of Science®Google Scholar
He W, Brumos J, Li H, Ji Y, Ke M, Gong X, Zeng Q, Li W, Zhang X, An F et al. 2011. A small-molecule screen identifies L-kynurenine as a competitive inhibitor of TAA1/TAR activity in ethylene-directed auxin biosynthesis and root growth in Arabidopsis. Plant Cell 23: 3944–3960.
10.1105/tpc.111.089029
CASPubMedWeb of Science®Google Scholar
Huang DW, Sherman BT, Lempicki RA. 2009. Systematic and integrative analysis of large gene lists using DAVID bioinformatics resources. Nature Protocols 4: 44–57.
10.1038/nprot.2008.211
CASPubMedWeb of Science®Google Scholar
Ishige F, Takaichi M, Foster R, Chua N-H, Oeda K. 1999. A G-box motif (GCCACGTGCC) tetramer confers high-level constitutive expression in dicot and monocot plants. The Plant Journal 18: 443–448.
10.1046/j.1365-313X.1999.00456.x
CASWeb of Science®Google Scholar
Izumi K, Nakagawa S, Kobayashi M, Oshio H, Sakurai A, Takahashi N. 1988. Levels of IAA, cytokinins, ABA and ethylene in rice plants as affected by a gibberellin biosynthesis inhibitor, uniconazole-P. Plant and Cell Physiology 29: 97–104.
CASWeb of Science®Google Scholar
Kärkönen A, Kuchitsu K. 2015. Reactive oxygen species in cell wall metabolism and development in plants. Phytochemistry 112: 22–32.
10.1016/j.phytochem.2014.09.016
CASPubMedWeb of Science®Google Scholar
Kim H, Park P-J, Hwang H-J, Lee S-Y, Man-Ho O, Kim S-G. 2006. Brassinosteroid signals control expression of the AXR3/IAA17 gene in the cross-talk point with auxin in root development. Bioscience, Biotechnology, and Biochemistry 70: 768–773.
10.1271/bbb.70.768
CASPubMedWeb of Science®Google Scholar
Knox K. 2003. AXR3 and SHY2 interact to regulate root hair development. Development 130: 5769–5777.
10.1242/dev.00659
CASPubMedWeb of Science®Google Scholar
Leyser HM, Ottoline FB, Pickett SD, Estelle M. 1996. Mutations in the AXR3 gene of Arabidopsis result in altered auxin response including ectopic expression from the SAUR-AC1 promoter. The Plant Journal 10: 403–413.
10.1046/j.1365-313x.1996.10030403.x
CASPubMedWeb of Science®Google Scholar
Leyser O. 2018. Auxin signaling. Plant Physiology 176: 465–479.
10.1104/pp.17.00765
CASPubMedWeb of Science®Google Scholar
Li G, Zhu C, Gan L, Ng D, Xia K. 2015. GA3 enhances root responsiveness to exogenous IAA by modulating auxin transport and signalling in Arabidopsis. Plant Cell Reports 34: 483–494.
10.1007/s00299-014-1728-y
CASPubMedWeb of Science®Google Scholar
Li H, Cheng Y, Murphy A, Hagen G, Guilfoyle TJ. 2009. Constitutive repression and activation of auxin signaling in Arabidopsis. Plant Physiology 149: 1277–1288.
10.1104/pp.108.129973
CASPubMedWeb of Science®Google Scholar
Li L, Verstraeten I, Roosjen M, Takahashi K, Rodriguez L, Merrin J, Chen J, Shabala L, Smet W, Ren H et al. 2021. Cell surface and intracellular auxin signalling for H⁺ fluxes in root growth. Nature 599: 273–277.
10.1038/s41586-021-04037-6
CASPubMedWeb of Science®Google Scholar
Lindsey BE, Rivero L, Calhoun CS, Grotewold E, Brkljacic J. 2017. Standardized method for high-throughput sterilization of Arabidopsis seeds. Journal of Visualized Experiments 17: 1–7.
Google Scholar
Ljung K, Hul AK, Kowalczyk M, Marchant A, Celenza J, Cohen JD, Sandberg G. 2002. Biosynthesis, conjugation, catabolism and homeostasis of indole-3-acetic acid in Arabidopsis thaliana. Plant Molecular Biology 50: 309–332.
10.1023/A:1016024017872
PubMedWeb of Science®Google Scholar
Lord SJ, Velle KB, Dyche Mullins R, Fritz-Laylin LK. 2020. SuperPlots: communicating reproducibility and variability in cell biology. Journal of Cell Biology 219: e202001064.
10.1083/jcb.202001064
CASPubMedWeb of Science®Google Scholar
Lucas M, Godin C, Jay-Allemand C, Laplaze L. 2008. Auxin fluxes in the root apex co-regulate gravitropism and lateral root initiation. Journal of Experimental Botany 59: 55–66.
10.1093/jxb/erm171
CASPubMedWeb of Science®Google Scholar
Mähönen AP, ten Tusscher K, Siligato R, Smetana O, Díaz-Triviño S, Salojärvi J, Wachsman G, Prasad K, Heidstra R, Scheres B. 2014. PLETHORA gradient formation mechanism separates Auxin responses. Nature 515: 125–129.
10.1038/nature13663
CASPubMedWeb of Science®Google Scholar
Majda M, Robert S. 2018. The role of auxin in cell wall expansion. International Journal of Molecular Sciences 19: 951.
10.3390/ijms19040951
PubMedWeb of Science®Google Scholar
Mhamdi A, Van Breusegem F. 2018. Reactive oxygen species in plant development. Development 145: 1–28.
10.1242/dev.164376
Web of Science®Google Scholar
Monshausen GB, Miller ND, Murphy AS, Gilroy S. 2011. Dynamics of auxin-dependent Ca²⁺ and PH signaling in root growth revealed by integrating high-resolution imaging with automated computer vision-based analysis. The Plant Journal 65: 309–318.
10.1111/j.1365-313X.2010.04423.x
CASPubMedWeb of Science®Google Scholar
Nishimura T, Nakano H, Hayashi K-i, Niwa C, Koshiba T. 2009. Differential downward stream of auxin synthesized at the tip has a key role in gravitropic curvature via TIR1/AFBs-mediated auxin signaling pathways. Plant and Cell Physiology 50: 1874–1885.
10.1093/pcp/pcp129
CASPubMedWeb of Science®Google Scholar
Ouellet F, Overvoorde PJ, Theologis A. 2001. IAA17/AXR3: biochemical insight into an auxin mutant phenotype. Plant Cell 13: 829–841.
10.1105/tpc.13.4.829
CASPubMedWeb of Science®Google Scholar
Overvoorde P, Fukaki H, Beeckman T. 2010. Auxin control of root development. Cold Spring Harbor Perspectives in Biology 2: a001537.
10.1101/cshperspect.a001537
CASPubMedWeb of Science®Google Scholar
Patro R, Duggal G, Love MI, Irizarry RA, Kingsford C. 2017. Salmon provides fast and bias-aware quantification of transcript expression. Nature Methods 14: 417–419.
10.1038/nmeth.4197
CASPubMedWeb of Science®Google Scholar
Pierre-Jerome E, Moss BL, Lanctot A, Hageman A, Nemhauser JL. 2016. Functional analysis of molecular interactions in synthetic auxin response circuits. Proceedings of the National Academy of Sciences, USA 113: 11354–11359.
10.1073/pnas.1604379113
CASPubMedWeb of Science®Google Scholar
Prerostova S, Dobrev PI, Knirsch V, Jarosova J, Gaudinova A, Zupkova B, Prášil IT, Janda T, Brzobohatý B, Skalák J et al. 2021. Light quality and intensity modulate cold acclimation in Arabidopsis. International Journal of Molecular Sciences 22: 2736.
10.3390/ijms22052736
CASPubMedWeb of Science®Google Scholar
Prigge MJ, Platre M, Kadakia N, Zhang Y, Greenham K, Szutu W, Pandey BK, Bhosale RA, Bennett MJ, Busch W et al. 2020. Genetic analysis of the arabidopsis TIR1/AFB auxin receptors reveals both overlapping and specialized functions. eLife 9: 1–28.
10.7554/eLife.54740
Web of Science®Google Scholar
Qi L, Kwiatkowski M, Chen H, Hoermayer L, Sinclair S, Zou M, del Genio CI, Kubeš MF, Napier R, Jaworski K et al. 2022. Adenylate cyclase activity of TIR1/AFB auxin receptors in plants. Nature 611: 133–138.
10.1038/s41586-022-05369-7
CASPubMedWeb of Science®Google Scholar
Rayle DL, Cleland RE. 1992. The acid growth theory of auxin-induced cell elongation is alive and well. Plant Physiology 99: 1271–1274.
10.1104/pp.99.4.1271
CASPubMedWeb of Science®Google Scholar
Reed JW. 2001. Roles and activities of Aux/IAA proteins in arabidopsis. Trends in Plant Science 6: 420–425.
10.1016/S1360-1385(01)02042-8
CASPubMedWeb of Science®Google Scholar
Rieu I, Eriksson S, Powers SJ, Gong F, Griffiths J, Woolley L, Benlloch R, Nilsson O, Thomas SG, Hedden P et al. 2008. Genetic analysis reveals that C19-GA 2-oxidation is a major gibberellin inactivation pathway in Arabidopsis. Plant Cell 20: 2420–2436.
10.1105/tpc.108.058818
CASPubMedWeb of Science®Google Scholar
Rizza A, Walia A, Lanquar V, Frommer WB. 2017. In vivo gibberellin gradients visualised in elongating tissues with gibberellin perception sensor. Nature Plants 10: 803–813.
10.1038/s41477-017-0021-9
Google Scholar
Růžička K, Ljung K, Vanneste S, Podhorská R, Beeckman T, Friml J, Benková E. 2007. Ethylene regulates root growth through effects on auxin biosynthesis and transport-dependent auxin distribution. Plant Cell 19: 2197–2212.
10.1105/tpc.107.052126
CASPubMedWeb of Science®Google Scholar
Sabatini S, Beis D, Wolkenfelt H, Murfett J, Guilfoyle T, Malamy J, Benfey P, Leyser O, Bechtold N, Weisbeek P et al. 1999. An auxin-dependent distal organizer of pattern and polarity in the Arabidopsis root. Cell 99: 463–472.
10.1016/S0092-8674(00)81535-4
CASPubMedWeb of Science®Google Scholar
Sarrion-Perdigones A, Falconi EE, Zandalinas SI, Juárez P, Fernández-del-Carmen A, Granell A, Orzaez D. 2011. GoldenBraid: an iterative cloning system for standardized assembly of reusable genetic modules. PLoS ONE 6: e21622.
10.1371/journal.pone.0021622
CASPubMedWeb of Science®Google Scholar
Sarrion-Perdigones A, Marta Vazquez-Vilar J, Palaci BC, Forment J, Ziarsolo P, Blanca J, Granell A, Orzaez D. 2013. GoldenBraid 2.0: a comprehensive DNA assembly framework for plant synthetic biology. Plant Physiology 162: 1618–1631.
10.1104/pp.113.217661
CASPubMedWeb of Science®Google Scholar
Sauer M, Robert S, Kleine-Vehn J. 2013. Auxin: simply complicated. Journal of Experimental Botany 64: 2565–2577.
10.1093/jxb/ert139
CASPubMedWeb of Science®Google Scholar
Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B et al. 2012. Fiji: an open-source platform for biological-image analysis. Nature Methods 9: 676–682.
10.1038/nmeth.2019
CASPubMedWeb of Science®Google Scholar
Schmitt J, Stunnenberg HG. 1993. The glucocorticoid receptor hormone binding domain mediates transcriptional activation in vitro in the absence of ligand. Nucleic Acids Research 21: 2673–2681.
10.1093/nar/21.11.2673
CASPubMedWeb of Science®Google Scholar
Serre NBC, Fendrych M. 2022. Acorba: automated workflow to measure Arabidopsis thaliana root tip angle dynamics. Quantitative Plant Biology 3: e9.
10.1017/qpb.2022.4
PubMedWeb of Science®Google Scholar
Serre NBC, Kralík D, Yun P, Slouka Z, Shabala S, Fendrych M. 2021. AFB1 controls rapid auxin signalling through membrane depolarization in Arabidopsis thaliana root. Nature Plants 7: 1229–1238.
10.1038/s41477-021-00969-z
CASPubMedWeb of Science®Google Scholar
Serre NBC, Wernerová D, Vittal P, Dubey SM, Medvecká E, Jelínková A, Petrášek J, Grossmann G, Fendrych M. 2023. The AUX1-AFB1-CNGC14 module establishes a longitudinal root surface PH profile. eLife 12: e85193.
10.7554/eLife.85193
CASPubMedWeb of Science®Google Scholar
Shani E, Weinstain R, Zhang Y, Castillejo C, Kaiserli E, Chory J, Tsien RY, Estelle M. 2013. Gibberellins accumulate in the elongating endodermal cells of Arabidopsis root. Proceedings of the National Academy of Sciences, USA 110: 4834–4839.
10.1073/pnas.1300436110
CASPubMedWeb of Science®Google Scholar
Sharma P, Jha AB, Dubey RS, Pessarakli M. 2012. Reactive oxygen species, oxidative damage, and antioxidative defense mechanism in plants under stressful conditions. Journal of Botany 2012: 1–26.
10.1155/2012/217037
CASGoogle Scholar
Shih H-W, DePew CL, Miller ND, Monshausen GB. 2015. The cyclic nucleotide-gated channel CNGC14 regulates root gravitropism in Arabidopsis thaliana. Current Biology 25: 3119–3125.
10.1016/j.cub.2015.10.025
CASPubMedWeb of Science®Google Scholar
Suzuki M, Yamazaki C, Mitsui M, Kakei Y, Mitani Y, Nakamura A, Ishii T, Soeno K, Shimada Y. 2015. Transcriptional feedback regulation of YUCCA genes in response to auxin levels in Arabidopsis. Plant Cell Reports 34: 1343–1352.
10.1007/s00299-015-1791-z
CASPubMedWeb of Science®Google Scholar
Swarup R, Kramer EM, Perry P, Kirsten Knox HM, Leyser O, Haseloff J, Beemster GTS, Bhalerao R, Bennett MJ. 2005. Root gravitropism requires lateral root cap and epidermal cells for transport and response to a mobile auxin signal. Nature Cell Biology 7: 1057–1065.
10.1038/ncb1316
CASPubMedWeb of Science®Google Scholar
Tanimoto E. 1987. Gibberellin-dependent root elongation in Lactuca sativa: recovery from growth retardant-suppressed elongation with thickening by low concentration of GA3. Plant and Cell Physiology 28: 963–973.
CASWeb of Science®Google Scholar
Tanimoto E. 2012. Tall or short? slender or thick? a plant strategy for regulating elongation growth of roots by low concentrations of gibberellin. Annals of Botany 110: 373–381.
10.1093/aob/mcs049
CASPubMedWeb of Science®Google Scholar
Tsukagoshi H, Busch W, Benfey PN. 2010. Transcriptional regulation of ROS controls transition from proliferation to differentiation in the root. Cell 143: 606–616.
10.1016/j.cell.2010.10.020
CASPubMedWeb of Science®Google Scholar
Ubeda-Tomás S, Swarup R, Coates J, Swarup K, Laplaze L, Beemster GTS, Hedden P, Bhalerao R, Bennett MJ. 2008. Root growth in Arabidopsis requires gibberellin/DELLA signalling in the endodermis. Nature Cell Biology 10: 625–628.
10.1038/ncb1726
CASPubMedWeb of Science®Google Scholar
Urfus T, Kopecký M, Urfusová R, Chrtek J. 2021. Whole genome duplication increases ecological niche breadth of the perennial herb Urtica dioica. Preslia 93: 305–319.
10.23855/preslia.2021.305
Web of Science®Google Scholar
Vanstraelen M, Benková E. 2012. Hormonal interactions in the regulation of plant development. Annual Review of Cell and Developmental Biology 28: 463–487.
10.1146/annurev-cellbio-101011-155741
CASPubMedWeb of Science®Google Scholar
von Wangenheim D, Hauschild R, Fendrych M, Barone V, Benková E, Friml J. 2017. Live tracking of moving samples in confocal microscopy for vertically grown roots. eLife 6: 26792.
10.7554/eLife.26792
PubMedWeb of Science®Google Scholar
Zhang H, Linster E, Gannon L, Leemhuis W, Rundle CA, Theodoulou FL, Wirtz M. 2019. Tandem fluorescent protein timers for noninvasive relative protein lifetime measurement in plants. Plant Physiology 180: 718–731.
10.1104/pp.19.00051
CASPubMedWeb of Science®Google Scholar
Zhao Y, Christensen SK, Fankhauser C, Cashman JR, Cohen JD, Weigel D, Chory J. 2001. A role for flavin monooxygenase-like enzymes in auxin biosynthesis. Science 291: 306–309.
10.1126/science.291.5502.306
CASPubMedWeb of Science®Google Scholar
Zuo J, Niu Q-W, Chua N-H. 2000. An estrogen receptor-based transactivator XVE mediates highly inducible gene expression in transgenic plants. The Plant Journal 24: 265–273.
10.1046/j.1365-313x.2000.00868.x
CASPubMedWeb of Science®Google Scholar

Citing Literature

Volume241, Issue6

March 2024

Pages 2448-2463

Auxin co-receptor IAA17/AXR3 controls cell elongation in Arabidopsis thaliana root solely by modulation of nuclear auxin pathway

Summary

Introduction

Materials and Methods