1. Developmental Biology
  2. Plant Biology
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Direct ETTIN-auxin interaction controls chromatin states in gynoecium development

  1. André Kuhn
  2. Sigurd Ramans Harborough
  3. Heather M McLaughlin
  4. Bhavani Natarajan
  5. Inge Verstraeten
  6. Jiří Friml
  7. Stefan Kepinski
  8. Lars Østergaard  Is a corresponding author
  1. Department of Crop Genetics, John Innes Centre, Norwich Research Park, United Kingdom
  2. Centre for Plant Sciences, Faculty of Biological Sciences, University of Leeds, United Kingdom
  3. Institute of Science and Technology, Austria
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Cite this article as: eLife 2020;9:e51787 doi: 10.7554/eLife.51787

Abstract

Hormonal signalling in animals often involves direct transcription factor-hormone interactions that modulate gene expression. In contrast, plant hormone signalling is most commonly based on de-repression via the degradation of transcriptional repressors. Recently, we uncovered a non-canonical signalling mechanism for the plant hormone auxin whereby auxin directly affects the activity of the atypical auxin response factor (ARF), ETTIN towards target genes without the requirement for protein degradation. Here we show that ETTIN directly binds auxin, leading to dissociation from co-repressor proteins of the TOPLESS/TOPLESS-RELATED family followed by histone acetylation and induction of gene expression. This mechanism is reminiscent of animal hormone signalling as it affects the activity towards regulation of target genes and provides the first example of a DNA-bound hormone receptor in plants. Whilst auxin affects canonical ARFs indirectly by facilitating degradation of Aux/IAA repressors, direct ETTIN-auxin interactions allow switching between repressive and de-repressive chromatin states in an instantly-reversible manner.

Introduction

Developmental programmes within multicellular organisms originate from a single cell (i.e. a fertilised oocyte) that proliferates into numerous cells ultimately differentiating to make up specialised tissues and organs. Tight temporal and spatial regulation of the genes involved in these processes is essential for proper development of the organism. Changes in gene expression are often controlled by mobile signals that translate positional information into cell type-specific transcriptional outputs (Hironaka and Morishita, 2012). In plants, this coordination can be facilitated by phytohormones such as auxin, which controls processes throughout plant development (Vanneste and Friml, 2009). In canonical auxin signalling, auxin-responsive genes are repressed when auxin levels are low by Aux/IAA transcriptional repressors that interact with DNA-bound Auxin Response Factors (ARFs). As auxin levels increase, the auxin molecule binds to members of the TIR1/AFB family of auxin co-receptors (Kepinski and Leyser, 2005; Dharmasiri et al., 2005). The main role of this auxin perception mechanism, besides non-transcriptionally inhibiting growth (Fendrych et al., 2018; Gallei et al., 2019), is transcriptional reprogramming. This is realised by interaction with Aux/IAA repressors, leading to their ubiquitination and subsequent degradation by the 26S proteasome, ultimately relieving the repression of ARF-targeted loci (Kelley and Estelle, 2012Leyser, 2018; Weijers and Wagner, 2016).

We recently identified an alternative auxin-signalling mechanism whereby auxin directly affects the activity of a transcription factor (TF) complex towards its downstream targets (Simonini et al., 2016; Simonini et al., 2017). This mechanism mediates precise polarity switches during organ initiation and patterning and includes the ARF, ETTIN (ETT/ARF3) as a pivotal component. However, ETT is an unusual ARF lacking the Aux/IAA-interacting Phox/Bem1 (PB1) domain (Simonini et al., 2016; Sessions et al., 1997) and is therefore unlikely to mediate auxin signalling via the canonical pathway. Here, we demonstrate that ETT binds auxin directly and that this interaction determines expression of ETT target genes. In conditions of low auxin levels, ETT interacts with co-repressors of the TOPLESS/TOPLESS-RELATED (TPL/TPR) family to keep chromatin at ETT target loci in a repressed state through HDA19-mediated histone deacetylation. Under high auxin conditions, ETT binds auxin leading to dissociation of TPL/TPR-HDA19 and de-repression of ETT targets.

Results

ETT can interact with a diverse set of TFs and these interactions are sensitive to the naturally occurring auxin, indole 3-acetic acid (IAA) (Simonini et al., 2016). To test if this effect is mediated through a direct interaction between ETT and auxin, we took advantage of the drug affinity-responsive target stability (DARTS) assay (Lomenick et al., 2011) and followed the effect of IAA on proteolytic degradation of a FLAG-fused ETT segment conditionally expressed in Arabidopsis (Figure 1a). The concentrations of IAA applied (0.1 µM, 1 µM, 10 µM) were selected to saturate the protein with ligand and to ensure maximal protection from proteolysis. As a negative control, benzoic acid (BA) was applied at the same dose as IAA (10 µM and 100 µM) because of its similar size and pKa as IAA. Treatments with IAA but not BA protected ETT-FLAG from pronase-induced degradation consistent with direct, specific interaction between IAA and ETT in planta.

Figure 1 with 2 supplements see all
ETT directly binds auxin (IAA).

(a) DARTS assay suggests that ETT binds IAA. Seedlings from an inducible two-component ETT-FLAG line were used for the protein isolation. Samples were treated with DMSO (mock), IAA and BA and digested by different concentrations of Pronase. Samples were further analysed by western blot with an anti-FLAG antibody. (b) HSQC-NMR performed with ES388-594 protein either alone (black), with indole-3-acetic acid (IAA, green) or benzoic acid (BA, pink). ES388-594: ligand, 1:10, 50 µM:500 µM. (c) Zoom-in of the indicated rectangular region in a. (d) Zoom-in of the specific shifts (labelled I-V) in the indicated dotted rectangles in b and c. Changes in chemical shifts are indicated by arrows from control to IAA treatment. (e–g) ITC spectre showing heat exchange between ES388-594 protein and IAA (e), but not in controls (f, g). See Figure 1—figure supplement 1 for parameters used in the HSQC-NMR experiment.

The region responsible for IAA-sensitivity is situated within the C-terminal part of ETT, known as the ETT-Specific (ES) domain. A protein fragment containing 207 amino acids of the ES domain, ES388-594, sufficient for mediating IAA-sensitivity in ETT-protein interactions, was produced recombinantly and shown to be intrinsically disordered (Simonini et al., 2018). The sensitivity of ETT-TF interactions to IAA suggests a direct effect of the IAA molecule on the ETT protein. Therefore, to test whether ETT binds IAA, we carried out heteronuclear single quantum coherence (HSQC) nuclear magnetic resonance (NMR) experiments using 15N-labelled ES388-594 protein. The HSQC spectrum, recorded at 5 °C, shows a prominent signal-dense region consistent with the ES domain being largely intrinsically disordered. Interestingly, the spectrum also shows dispersed peaks flanking the signal-dense region indicating that there is nevertheless some propensity to form secondary structure, particularly with a helical character (Figure 1b). In addition to this overview of ETT structure, the HSQC NMR probes chemical shifts of protein amide-NH bonds in response to the presence of ligand (Meyer and Peters, 2003). We found that a number of residues shifted their position in the spectrum in response to the addition of IAA, whereas addition of BA had no effect (Figure 1b–d). These shifts show that certain residues are experiencing a changed chemical environment as a consequence of IAA-binding and this may include the conformational change of a structural motif within the ETT protein. The HSQC experiment therefore demonstrates that ETT binds IAA directly. This experiment has not allowed us to assign signals to specific amino acids and hence there is some uncertainty associated with tracking the chemical shifts of some residues. However, a particularly large change is observed for the tryptophan NH cross peak when IAA is added to the ETT fragment (~10 ppm, rectangle I in Figure 1b,d). Since there is only one tryptophan in the ETT fragment used here (W505), this shift can be assigned to this residue and suggests that W505 has a prominent role in the interaction with IAA.

We also used the recombinant ETT fragment in an Isothermal Titration Calorimetry (ITC) assay, which characterises binding of ligands to proteins by determining thermodynamic parameters of the interaction as heat exchange. This experiment revealed interaction between ETT and IAA, while control experiments titrating IAA into buffer without protein and titrating buffer without IAA into the ETT fragment showed no heat exchange (Figure 1e–g). We were unable to obtain a Kd value for the interaction, which may be due to the interaction being weak as well as potential protein aggregation. This could reflect that the ETT-auxin interaction is stabilised by interacting protein partners in planta that are not present in this in vitro experiment.

Together, these three independent biochemical methods demonstrate that ETT binds IAA directly thus revealing a key molecular aspect of the non-canonical auxin-signalling pathway.

Previously, PINOID (PID) (Benjamins et al., 2001) and HECATE1 (HEC1) (Gremski et al., 2007) were identified as ETT target genes (Simonini et al., 2016; Simonini et al., 2017) and both genes are upregulated in gynoecium tissue from the ett-3 mutant compared to wild type (Figure 2—figure supplement 1). We also observed that expression of both genes is induced by IAA, but did not observe any additional induction beyond the constitutive upregulation in the ett-3 mutant background (Figure 2—figure supplement 1). This ETT-dependent regulation does not require a functional TIR1/AFB machinery, since IAA-induction of PID and HEC1 is still observed in tir1/afb mutant combinations, whereas the known TIR1/AFB-mediated auxin induction of the IAA19 gene is completely abolished in these mutants (Figure 2a–c).

Figure 2 with 1 supplement see all
ETT regulates target gene expression independently of TIR1/AFB auxin receptors.

Expression of the canonical auxin responsive IAA19 gene (a) and the ETT-target genes HEC1 (b) and PID (c) in control-treated (dark grey) or 100 µM IAA-treated (light grey) gynoecia assayed using qRT-PCR. (a) IAA19 expression is up-regulated in response to auxin in wild-type gynoecia (Col-0) but not in tir1/afb double and triple mutants. The ETT-target genes HEC1 and PID are up-regulated in response to auxin in both wild-type and auxin receptor mutants (b c). This suggests a TIR1/AFB independent regulation of these genes. (d) Expression of IAA19, HEC1 and PID in response to treatment with 100 µM IAA and 100 µM cvxIAA in wild-type (Col-0) and pTIR1:ccvTIR1 gynoecia in the tir1 afb2 double mutant (ccvTIR1). The data confirm TIR1/AFB independent regulation of HEC1 and PID in the gynoecium. ***p<0.0001; Shown are mean ± standard deviation of three biological replicates. See Figure 2—source data 2 for statistical analyses.

To further assess the TIR1/AFB independence of the ETT-mediated auxin signalling pathway, we exploited a recently-developed synthetic auxin-TIR1 pair (Uchida et al., 2018). In this system, the auxin-binding pocket of TIR1 has been engineered (ccvTIR1) to accommodate an IAA derivative bearing a bulky side chain (cvxIAA). By expressing the ccvTIR1 in a tir1 afb2 mutant background, the canonical pathway will only respond to the addition of cvxIAA and not IAA (Uchida et al., 2018). We performed an expression analysis on ccvTIR1 gynoecia treated ±cvxIAA and ±IAA as well as control plants with the same treatments. In this experiment, IAA19 served as a control gene whose expression is regulated in a TIR1/AFB-dependent manner (Benjamins et al., 2001). Indeed, IAA19 was strongly upregulated by cvxIAA in the ccvTIR1 line, but not by IAA (Figure 2d). In contrast, PID and HEC1 expression was not significantly affected by cvxIAA, whilst still responding to IAA in the ccvTIR1 background (Figure 2d). These data demonstrate that ETT-mediated auxin signalling can occur independently of the canonical TIR1/AFB signalling pathway.

In a phylogenetic analysis of ETT protein sequences across the angiosperm phylum, we identified a number of regions that are highly conserved (Figure 3—figure supplement 1). Unsurprisingly, the DNA-binding domain characteristic to B3-type TFs such as ARF proteins was conserved across all ETT proteins. Towards the C terminus of the ES domain we identified an EAR-like motif with a particularly high level of conservation (Figure 3a, Figure 3—figure supplement 1a). Ethylene-responsive element binding factor-associated Amphiphilic Repression (EAR) motifs are also found in Aux/IAA proteins. Interactions between Aux/IAA and members of the TOPLESS and TOPLESS-RELATED (TPL/TPR) family of co-repressors occur via this motif (Szemenyei et al., 2008). TPL/TPRs mediate their repressive effect by attracting histone deacetylases (HDACs) to promote chromatin condensation (Krogan et al., 2012). It was recently shown that ETT is required to keep Histone H3 at a deacetylated state at the gene locus of the meristem identity gene SHOOT MERISTEMLESS (STM) thereby repressing STM expression (Chung et al., 2019). Since ETT functions independently of the canonical auxin pathway, it is possible that its role in chromatin remodelling occurs via direct interaction with TPL/TPRs through the EAR-like motif. To test this, we carried out Yeast 2-Hybrid (Y2H) assays in which ETT was found to interact with TPL, TPR2 and TPR4 (Figure 3b, Figure 3—figure supplement 1b). Moreover, mutating residues in the EAR-like motif abolished the interactions demonstrating its requirement for the ETT-TPL/TPR interaction (Figure 3b).

Figure 3 with 3 supplements see all
ETT interacts with members of the TPL/TPR co-repressor family in an auxin-sensitive manner.

(a) Schematic representation of full-length ETT protein highlighting an EAR-like motif and tryptophan in position 505 (W) in the C-terminal ETT-specific domain. DBD, DNA-binding domain. (b) Y2H showing that ETT interacts with TPL, TPR2 and TPR4. These interactions depend on the identified C-terminal RLFGF motif and are auxin-sensitive. DBD, DNA-binding domain. (c) Co-IP revealing that ETT interacts with TPL in an auxin-sensitive manner with increasing IAA concentrations weakening the interaction. (d) Co-IP showing that mutating the tryptophan in position 505 of ETT (ETTW505A) leads to loss of auxin sensitivity in the interaction between ETT and TPL.

Given that several ETT-protein interactions are affected by IAA and that part of the ETT transcriptome changes in response to IAA (Simonini et al., 2016; Simonini et al., 2017), we tested the IAA sensitivity of ETT-TPL/TPR interactions. In both Y2H and in co-immunoprecipitation (Co-IP) experiments, we observed that the interactions were reduced with increasing IAA concentrations (Figure 3b,c and Figure 3—figure supplement 2). Moreover, as described previously for other ETT-protein interactions, the sensitivity was specific to IAA as other auxinic compounds tested did not show this effect (Figure 3—figure supplement 2). Henceforth, ‘auxin’ will refer to IAA unless stated otherwise. These data suggest that in conditions with low auxin levels, ETT can interact with TPL/TPR proteins to repress the expression of target genes. An increase in cellular auxin causes ETT to bind auxin thereby undergoing a conformational change that abolishes interaction with TPL/TPR co-repressors.

The HSQC-NMR experiment described above indicate that the tryptophan in position 505 (W505) affects the direct interaction between the ETT protein and auxin. To test the role of W505 in the auxin-sensitive interaction with TPL/TPRs, we constructed a mutated version of ETT (W505A) and assessed the effect in protein interaction assays. Both Y2H and Co-IP experiments revealed that W505 is required for the ETT-TPL/TPR interaction to be sensitive to auxin (Figure 3c,d). The key importance of W505 is furthermore supported by a phylogenetic analysis of ETT protein sequences revealing that this residue is highly conserved in ETT proteins across angiosperms (Figure 3—figure supplement 1a).

TPL was originally identified as a key factor involved in setting up the apical-basal growth axis during embryo development (Long et al., 2006; Smith and Long, 2010). Large-scale interaction studies suggest that the five Arabidopsis TPL/TPRs have roles throughout plant development (Krogan et al., 2012; Causier et al., 2012). Whilst ETT has been implicated in a wide array of developmental processes (Garcia et al., 2006; Marin et al., 2010; Kelley et al., 2012; Pekker et al., 2005), the most dramatic phenotypes of ett loss-of-function mutants are observed during gynoecium development (Sessions et al., 1997; Sessions and Zambryski, 1995; Nemhauser et al., 2000). In accordance with this, ETT is highly expressed in the gynoecium (Figure 4a; Simonini et al., 2016). We produced reporter lines of TPL, TPR2 and TPR4 promoters fused to the GUS gene to test if they overlap with ETT expression in the gynoecium. Both pTPL:GUS and pTPR2:GUS exhibited strong expression in the apical part of the gynoecium where ETT is also expressed, while no pTPR4:GUS expression was observed (Figure 4a–d). Single loss-of-function mutants in TPL and TPR2 do not show any abnormal phenotypes during gynoecium development. However, the tpl tpr2 double mutant has defects in the development of the apical gynoecium similar to ett mutants (Figure 4e–g) demonstrating that TPL and TPR2 function redundantly in gynoecium development. Together with the protein interaction data and the overlapping expression patterns, these results suggest that ETT and TPL/TPR2 cooperate to regulate gynoecium development.

Figure 4 with 1 supplement see all
ETT, TPL/TPR2 and HDA19 co-operatively regulate gene expression to facilitate gynoecium development.

(a–d) Promoter GUS expression analysis of pETT:GUS (a), pTPL:GUS (b), pTPR2:GUS (c) and pTPR4:GUS (d) revealed that ETT, TPL and TPR2 but not TPR4 are co-expressed in the Arabidopsis style. Scale bar = 300 μm. (e–h) Gynoecium phenotypes of wild-type (e), ett-3 (f) tpl tpr2ge (g) and hda19-4 (h). Scale bar = 100 μm. (i j,) HEC1 (i) and PID (j) are constitutively mis-regulated in ett-3, tpl tpr2ge and hda19-4 gynoecia. This misregulation is unaffected by treatment with 100 µM IAA. ***p-values<0.0001; Shown are mean ± standard deviation of three biological replicates. Differences between untreated and IAA-treated mutants are not significant. See Figure 4—source data 1 for statistical analyses.

TPL was shown previously to recruit histone deacetylase, HDA19, during early Arabidopsis flower development to keep chromatin in a repressed state (Krogan et al., 2012). Moreover, HDA19 was also recently shown to participate in the repression of STM (Chung et al., 2019). In that study, ETT was shown to recruit HDA19 to the STM promoter, although not via direct protein interaction. Here, our analysis of gynoecia from the hda19-4 mutant demonstrate that HDA19 is also required for gynoecium development as the hda19-4 mutant has strong style defects (Figure 4h). In agreement with this, the HDA19 gene was highly expressed in gynoecium tissue, whereas another member of the HDA gene family, HDA6, was not (Figure 4—figure supplement 1). Moreover, HDA19 recruitment likely involves ETT, since expression of the ETT target genes, PID and HEC1, is increased in the tpl tpr2 and hda19-4 mutants compared to wild type. Similar to the ett mutant, auxin treatments failed to further induce expression in these mutants (Figure 4i,j). These observations suggest that ETT, TPL/TPR2 and HDA19 function in conjunction to control gene expression during gynoecium development.

To test the direct interaction of ETT, TPL and HDA19 on chromatin, we performed Chromatin-Immunoprecipitation (ChIP) using reporter lines expressing GFP fusion protein. Although only ETT is expected to bind DNA, ChIP followed by qPCR revealed that all three proteins associate with DNA elements in the same regions of the promoters of PID and HEC1 (Figure 5a–c). Moreover, while ETT association with these promoter regions was largely unaffected in the presence of IAA, TPL and HDA19 interactions are reduced upon auxin treatment (Figure 5a–c). This supports a model in which ETT recruits TPL/TPR2 and HDA19 to ETT target loci to keep chromatin in a condensed state through histone deacetylation. When auxin levels increase, the ETT-TPL/TPR2 interaction is broken and TPL/TPR2 and HDA19 dissociate from the loci, presumably preventing HDA19 from deacetylating histones. To test this, we assayed for H3K27 acetylation, which is a substrate for HDA19. H3K27 acetylation increased in the absence of ETT and upon treatment with auxin. This occurred in the same regions of the PID and HEC1 promoters where the proteins were found to associate (Figure 5d,e). In agreement with ETT mediating the association of TPL/TPR and HDA19 with these regions, there was no further increase of acetylation in the ett-3 mutant upon treatment with auxin (Figure 5d,e).

ETT, TPL and HDA19 co-operatively regulate HEC1 and PID by modulating chromatin acetylation.

(a–c) Chromatin immunoprecipitation (ChIP) shows ETT (a), TPL (b) and HDA19 (c) binding to conserved regions of HEC1 and PID loci in the absence of IAA (dark grey columns). In the presence of IAA (light grey columns), ETT association is unchanged, while both TPL and HDA19 interactions are reduced. A WUS gene fragment served as negative control (NC). (d e,) H3K27ac accumulation (from ChIP analysis) along the HEC1 (d) and PID (e) loci in wild-type (Col-0) and ett-3 plants ± treatment with 100 µM IAA. Numbers on the x axes are distances to the Transcription Start Site (TSS). The schematic of the loci is shown below each panel. Dashed boxes represent ETT binding regions. (f) Qualitative schematic illustration of alternative TIR1/AFB independent auxin signaling. Under low auxin conditions an ETT-TPL-HDA19 complex binds to ETT-target genes keeping their chromatin environments repressed, through de-acetylation. High nuclear auxin concentrations abolish the ETT-TPL-HDA19 complex through direct ETT-auxin interaction. This leads to an accumulation of histone acetylation and up-regulation of ETT-target genes. Values in a–e) are means ± standard deviation of three biological replicates. See Figure 5—source data 1 for statistical analyses.

Discussion

The data presented in this paper provide molecular insight into how auxin levels are translated into changes in gene expression of ETT target genes. Our data lead to a model in which low levels of auxin maintain ETT associations with TPL/TPR2 to repress gene expression via H3K27 deacetylation. As auxin levels increase, TPL/TPR2 (and hence HDA19) disassociate from ETT, promoting H3K27 acetylation (Figure 5f). This model molecularly underpins the published association between auxin dynamics and PID expression at the gynoecium apex where PID is repressed at early stages of development to allow symmetry transition, but subsequently de-repressed as auxin levels rise to facilitate polar auxin transport (Simonini et al., 2016; Moubayidin and Ostergaard, 2014). Whilst this model explains the data on ETT-mediated auxin signalling during gynoecium development, it is possible that the effect of auxin on ETT varies depending on the developmental context. Moreover, the concentrations of auxin required to mediate its effect on ETT is likely to differ between diverse ETT-containing complexes.

The direct binding of auxin allows ETT to switch the chromatin locally between repressive and de-repressive states. The recent report of ETT negatively regulating STM expression (Chung et al., 2019) provides molecular evidence that ETT can function as a repressor. Whether ETT also has a role in recruiting activating components is yet to be clarified. Indeed, different modes of actions are possible that could depend on other interacting partners or auxin levels and even be target-gene specific. Nevertheless, the effect of auxin on ETT-mediated regulation of the genes studied here is instantly reversible, making it possible to switch between states, immediately responding to changes in auxin levels. This feature, which is reminiscent of animal hormonal signalling pathways such as the Thyroid Hormone and Wnt/ß-catenin pathways (Tsai and O'Malley, 1994; Gammons and Bienz, 2018), may be particularly important in controlling changes in tissue polarity during plant organogenesis as observed in the Arabidopsis gynoecium (Moubayidin and Ostergaard, 2014).

The identification of a direct auxin-ETT interaction to control gene expression adds an additional layer of complexity to auxin action, which contributes towards explaining how auxin imparts its effect on highly diverse processes throughout plant development. In a broader context, this work also opens for the exciting possibility that direct transcription factor-ligand interactions is a general feature in the control of gene expression in plants as found in animals.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional
information
Gene (Arabidopsis thaliana)ETTIN (ETT)The Arabidopsis Information ResourceAT2G33860
Gene (Arabidopsis thaliana)TOPLESS
(TPL)
The Arabidopsis Information ResourceAT1G15750
Gene (Arabidopsis thaliana)TOPLESS- RELATED 1 (TPR1)The Arabidopsis Information ResourceAT1G80490
Gene
(Arabidopsis thaliana)
TOPLESS-RELATED 2 (TPR2)The Arabidopsis Information ResourceAT3G16830
Gene (Arabidopsis thaliana)TOPLESS-RELATED 3 (TPR3)The Arabidopsis Information ResourceAT5G27030
Gene (Arabidopsis thaliana)TOPLESS-RELATED 4 (TPR4)The Arabidopsis Information ResourceAT3G15880
Gene (Arabidopsis thaliana)HISTONE DEACETYLASE 6
(HDA6)
The Arabidopsis Information ResourceAT5G63110
Gene (Arabidopsis thaliana)HISTONE DEACETYLASE 19
(HDA19)
The Arabidopsis Information ResourceAT4G38130
Gene (Arabidopsis thaliana)HECATE 1
(HEC1)
The Arabidopsis Information ResourceAT5G67060
Gene (Arabidopsis thaliana)PINOID (PID)The Arabidopsis Information ResourceAT2G34650
Genetic reagent Arabidopsis thalianaCol-0widely distributed
Genetic reagent Arabidopsis thalianahda19-4Kim et al., 2008SALK_13944
Genetic reagent Arabidopsis thalianatir1-1 afb2-3 afb3-4Parry et al., 2009
Genetic reagent Arabidopsis thalianatplEuropean Arabidopsis Stock CentreSALK_034518C
Genetic reagent Arabidopsis thalianaett-3Simonini et al., 2017Sessions et al., 1997AT2G33860
Genetic reagent Arabidopsis thalianapETT:GUSNg et al., 2009AT2G33860
Genetic reagent Arabidopsis thalianapETT:ETT-GFPSimonini et al., 2016AT2G33860
Genetic reagent Arabidopsis thaliana pTPL:TPL-GFPPi et al., 2015AT1G15750
Genetic reagent Arabidopsis thalianap35S:HDA19-GFPPi et al., 2015AT4G38130
Genetic reagent Arabidopsis thalianapTIR1:ccvTIR1Szemenyei et al., 2008
Genetic reagent Arabidopsis thaliana tpl tpr2gethis paperAT1G15750, AT3G16830Further details in the Materials and methods section
Genetic reagent Arabidopsis thalianapTPL:GUSthis paperAT1G15750Further details in the Materials and methods section
Genetic reagent Arabidopsis thalianapTPR2:GUSthis paperAT3G16830Further details in the Materials and methods section
Genetic reagent Arabidopsis thalianapTPR4:GUSthis paperAT3G15880Further details in the Materials and methods section
Genetic reagent Arabidopsis thalianap35S:CDGVG p6xGAL4UAS:ETT-FLAGthis paperFurther details in the Materials and methods section
AntibodyMouse anti-FLAGAbcamab49763monoclonal, conjugated with HRP
AntibodyMouse anti-HAAbcamab173826monoclonal, conjugated with HRP
AntibodyMouse anti-GFPRoche11814460001monoclonal
AntibodyRabbit anti-H3K27acAbcamab4729polyclonal
AntibodyRabbit anti-H3Abcamab1791polyclonal
Recombinant DNA reagentpGWB14; p35s:HA-TPLEspinosa-Ruiz et al., 2017
Recombinant DNA reagentpICH47732; p35s:ETT-3xFLAGthis paperBackbone Addgene
#48000
Recombinant DNA reagentpICH47732; p35s:
ETTW505A-3xFLAG
this paperBackbone Addgene
#48000
Recombinant DNA reagentpGADT7; ETTthis paperFurther details in the Materials and methods section
Recombinant DNA reagentpGADT7; ETTW505Athis paperFurther details in the Materials and methods section
Recombinant DNA reagentpGADT7; ETTL552S; F553Sthis paperFurther details in the Materials and methods section
Recombinant DNA reagentpGBKT7; TPLCausier et al., 2012
Recombinant DNA reagentpGBKT7; TPR1Causier et al., 2012
Recombinant DNA reagentpGBKT7; TPR2Causier et al., 2012
Recombinant DNA reagentpGBKT7; TPR3Causier et al., 2012
Recombinant DNA reagentpGBKT7; TPR4Causier et al., 2012
Recombinant DNA reagentpESPRIT; ES388-594Simonini et al., 2018

Plant materials and treatments

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Plants were grown in soil at 22°C in long day conditions (16 hrs day/8 hr dark). All mutations were in the Col-0 background. Mutant alleles described before include ett-3 (Sessions et al., 1997; Simonini et al., 2016), hda19-4 (SALK_139443) (Kim et al., 2008), pETT:GUS (Ng et al., 2009), pETT:ETT-GFP in ett-3 (Simonini et al., 2016), pTPL:TPL:GFP (Pi et al., 2015), p35S:HDA19:GFP (Pi et al., 2015), pTIR1:ccvTIR1 in tir1-1 afb2-3 (Szemenyei et al., 2008) and tir1-1 afb2-3 afb3-4(Parry et al., 2009). The tpl mutant (SALK_034518C) was obtained from the European Arabidopsis Stock Centre.

For both expression and ChIP analysis, auxin treatments were applied by spraying bolting Col-0 and ett-3 inflorescences with a solution containing 100 µM IAA (Sigma) or cvxIAA and 0.015% Silwet L-77 (De Sangosse Ltd.). Treated samples were returned to the growth room and incubated for two hours.

Expression analysis

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Quantitative Real time PCR (qRT-PCR) was used for expression analysis. RNA was extracted from floral buds using the RNeasy mini kit (Qiagen). Using the SuperScript IV First-Strand Synthesis kit (ThermoFisher), cDNA was synthesised from 1 µg of total RNA. Subsequently, qRT-PCR was carried out using SYBR Green JumpStart Taq ReadyMix (Sigma) using the appropriate primers (Figure 2—source data 1). Relative expression values were determined using the 2-ΔΔCt method (Livak and Schmittgen, 2001). Data were normalised to POLYUBIQUITIN 10 (UBQ10/AT4G05320) expression.

ETT protein analysis by alignment

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Published ETT sequences of 22 Angiosperm species were retrieved from Phytozome version 12 (Goodstein et al., 2012). Nucleotide sequences were translated and aligned using MUSCLE in Geneious version 6.1.8 (Kearse et al., 2012). The EAR-like motif was extracted as a sequence logo (Figure 3a; Figure 3—figure supplement 1).

Generation of the tpl tpr2 CRISPR mutant

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The tpl tpr2ge mutant was generated using CRISPR/Cas9 technology by a method previously described (Castel et al., 2019). Briefly, for the construction of the RNA-guided genome-editing plasmid, DNA sequences encoding the gRNA adjacent to the PAM sequences were designed to target two specific sites in TPR2 (AT3G16830). DNA-oligonucleotides (Figure 2—source data 1) containing the specific gRNA sequence were synthesised and used to amplify the full gRNA from a template plasmid (AddGene #46966). Using Golden Gate cloning (Engler et al., 2014) each gRNA was then recombined in a L1 vector downstream of U6 promoter (Goodstein et al., 2012). Finally, the resulting gRNA plasmids were then recombined with a L1 construct containing pYAO:Cas9_3:E9t (Castel et al., 2019) (kindly provided by Jonathan Jones) and a L1 construct containing Fast-Red selection marker (AddGene #117499) into a L2 binary vector (AddGene #112207).

The construct was transformed into Agrobacterium tumefaciens strain GV3101 by electroporation, followed by plant transformation by floral dip into the tpl single mutant (Clough and Bent, 1998). Transgenic T1 seeds appear red under UV light and were selected under a Leica M205FA stereo microscope. T1 plants were genotyped using PCR and the TPR2 locus sequenced (Oligonucleotides in Figure 2—source data 1). Genome-edited plants were selected and the next generation grown (T2). Seeds of this generation were segregating in a 3:1 ratio for the transgene. Transgene negative plants were selected and grown on soil. To identify homozygous mutations T2 plants were genotyped. The T3 generation was again checked for the absence of the transgene.

Protein interaction

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For Yeast-two-Hybrid (Y2H) assays coding sequences were cloned into pDONR207 and recombined into the pGDAT7 and pGBKT7 (Clontech). Using the co-transformation techniques (Engler et al., 2014) these constructs were transformed into the AH109 strain (Clontech). Transformations were selected on Yeast Selection Medium (YSD) lacking Tryptophan (W) and Leucine (L) at 28°C for 3–4 days. Transformed yeast cells were serially diluted (100, 10-1, 10-2 and 10-3) and dotted on YSD medium lacking Tryptophan (W), Leucin (L), adenine (A) and Histidine (H) to test for interaction. To examine interaction strength 3-amino-1,2,4-triazole (3-AT) was supplemented to the YSD (-W-L-A-H) medium with different concentrations (0, 5, 10 mM). To determine the effect of auxinic compounds on the protein-protein interactions benzoic acid (BA), IAA, NAA and 2,4D (all Sigma) were dissolved in ethanol and added directly to the medium at the desired concentrations. Pictures were taken after 3 days of growth at 28°C.

For the β-Galactosidase assay, transgenic yeast was grown in liquid YSD (-W-L) medium supplemented with /-out 100 µM IAA or NAA, to an OD600 of 0.5. The cells were then harvested and lysed using 150 µL Buffer Z with β-mercaptoethanol (100 mM Phosphate buffer pH 7, 10 mM KCl, 1 mM Mg2SO4, β-mercaptoethanol 50 mM), 50 µL chloroform and 20 µL of 0.1% SDS. After lysis, the sample was incubated with 700 µL pre-warmed ONPG solution (1 mg/mL ONPG (o-Nitrophenyl-β-D-Galactopyranoside, Sigma) prepared in Buffer Z without β-mercaptoethanol at 37°C until a yellow colour developed in the samples without auxin treatment. After stopping all reactions (using 500 µL Na2CO3) the supernatant was collected and OD405 determined. The β-Galactosidase activity was calculated as follows: (A405*1000)/(A600*min*mL).

For co-immunoprecipitation, ETT-FLAG was generated using Golden Gate cloning (Kearse et al., 2012) by recombining a previously described L0 clone for ETT (Simonini et al., 2016) with a 35S promoter (AddGene #50266), a C-terminal 3xFLAG epitope (AddGene #50308) and a Nos-terminator (AddGene #50266) into a L1 vector (AddGene #48000). The pGWB14 TPL-HA construct was provided by Salomé Prat and has been used in previous studies (Espinosa-Ruiz et al., 2017). The epitope-tagged proteins were transiently expressed in four-week-old N. benthamiana leaves for two days. Co-immunoprecipitation was performed as described previously (Egea-Cortines et al., 1999). After harvest, 1 g of fresh leaf tissue was ground in liquid nitrogen. The powder was homogenised for 30 min in two volumes of extraction buffer (10% glycerol, 25 mM Tris-HCl pH 7.5, 1 mM EDTA, 150 mM NaCl, 0.15% NP-40, 1 mM PMSF, 10 mM DTT, 2% Polyvinylporrolidone, 1x cOmplete Mini tablets EDTA-free Protease Inhibitor Cocktail (Roche). The homogenised samples were cleared by centrifugation at 14,000 x g for 10 min and cleared lysates were incubated for 2 hr with 20 µl anti-FLAG M2 magnetic beads (SIGMA-ALDRICH, M8823; lot: SLB2419). The beads were washed five times with IP buffer (10% glycerol, 25 mM Tris-HCl pH 7.5, 1 mM EDTA, 150 mM NaCl, 0.15% NP-40, 1 mM PMSF, 1 mM DTT, 1x cOmplete Mini tablets EDTA-free Protease Inhibitor Cocktail (Roche)) and proteins were eluted by adding 80 µl 2x SDS loading buffer followed by an incubation at 95°C for 10 min. To examine auxin sensitivity, 4 g of fresh leaf tissue was collected, ground in liquid nitrogen and protein was extracted. The lysate was then divided according to the number of treatments. The desired concentration of IAA or NAA was added to each of the cleared lysates before the anti-FLAG M2 magnetic beads were added. IAA or NAA at the desired concentration was also supplemented to the IP buffer during the washes. The eluates were analysed by western blot using an anti-FLAG antibody (M2, Abcam, ab49763, Lot: GR3207401-3) or an anti-HA antibody (Abcam, ab173826, Lot: GR3255539-1). Both antibodies were used as 1:10000 dilutions. The antibodies were validated by the manufacturer.

Scanning electron microscopy

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Whole inflorescences of Col-0, ett-3, tpl tpr2ge and hda19-4 were fixed overnight in FAA (3.7% formaldehyde, 5% glacial acetic acid, 50% ethanol) and dehydrated through an ethanol series (70% to 100%) as described previously (Moubayidin and Ostergaard, 2014). The samples were then critical point-dried, gynoecia dissected and mounted. After gold coating, samples were examined with a Zeiss Supra 55VP Field Emission Scanning electron microscope using an acceleration voltage of 3 kV.

TPL, TPR2 and TPR4 reporter lines

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For the construction of the promoter:GUS reporter plasmids of TPL, TPR2 and TPR4, 2.5 kb of promoter sequences were isolated from genomic DNA and inserted upstream of the ß-glucoronidase gene of pCambia1301 vectors using the In-Fusion Cloning Recombinase kit (Clontech). The constructs were transformed into Agrobacterium tumefaciens strain GV3101 by electroporation, followed by plant transformation by floral dip into Col-0 (Clough and Bent, 1998).

The GUS histochemical assay was performed in at least three individual lines per construct. Inflorescences of each GUS line were pre-treated with ice cold acetone for 1 hr at −20°C and washed two times for 5 min with 100 mM sodium phosphate buffer followed by one wash with sodium phosphate buffer containing 1 mM K3Fe(CN)6 and 1 mM K4Fe(CN)6 (both Sigma) at room temperature. Subsequently, samples were vacuum infiltrated for 5 min with X-Gluc solution (100 mM sodium phosphate buffer, 10 mM EDTA, 0.5 mM K3Fe(CN)6, 3 mM K4Fe(CN)6, 0.1% Triton X100) containing 1 mg/ml of ß-glucoronidase substrate X-gluc (5-bromo-4-chloro-3-indolylglucuronide, Melford) and incubated at 37°C. pTPL:GUS samples were incubated for 20 min and pTPR2:GUS lines for 45 min to prevent overstaining. pTPR4:GUS lines were incubated for 16 hr. After staining, the samples were washed in 70% ethanol until chlorophyll was completely removed. Gynoecia were dissected and mounted in chloral hydrate (Sigma). Samples were analysed using a Leica DM6000 light microscope.

Chromatin immunoprecipitation

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Transcription factor ChIP was performed in biological triplicates using the pETT:ETT:GFP, pTPL:TPL:GFP and p35S:HDA19-GFP lines and data analysed as described previously (Chung et al., 2019). Additionally, a WUS gene fragment (WUSp4 amplicon) was used as a negative control for ETT binding (Liu et al., 2014). IP was conducted using the anti-GFP antibody (Roche, 11814460001, Lot: 19958500) and Pierce Protein G magnetic beads (ThermoFisher, 88847, Lot: SI253639) were used for IP.

Histone acetylation ChIP was carried out and data were analysed as described previously (Qüesta et al., 2016). The experiment was carried out in triplicate using 3 g auxin-treated or untreated Col-0 or ett-3 inflorescent tissue. The antibodies used for IP were anti-H3K27ac antibodies (Abcam, ab4729, Lot: GR3231937-1) and anti-H3 (Abcam, ab1791, Lot: GR310541-1). All antibodies were validated by the manufacturers.

In all ChIP experiments, DNA enrichment was quantified using quantitative PCR (qPCR) with the appropriate primers (Figure 2—source data 1). In case of H3K27ac, H3 was used as an internal control and the data represented as ratio of (H3K27ac at HEC1 or PID divided by H3 at HEC1 or PID).

Statistical analyses and replication

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In all graphs error bars represent the standard deviation of the mean for all numerical values. QRT-PCR and ChIP experiments have been carried out at least in triplicate. The data presented here show an average of three replicates. For qRT-PCR data were analysed using one-way ANOVA with post-hoc Tukey multiple comparison test. ChIPqPCR data were analysed using two-way ANOVA with post hoc Bonferroni multiple comparison test. All output of statistical tests can be found in the source data files. All statistical tests were carried out using GraphPad Prism Version 5.04 (La Jolla California USA, www.graphpad.com).

Drug Affinity Responsive Target Stability (DARTS) assay

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The DARTS assay to evaluate the stability of ETTIN-FLAG in the presence of IAA was performed as described previously (Lomenick et al., 2011; Kania et al., 2018; Tan et al., 2020) with plant material of a two-component inducible ETTIN-FLAG line. This ETT-FLAG line was generated using Golden Gate cloning (Engler et al., 2014) by recombining a previously described L0 clone for ETT (Simonini et al., 2016) with a 6xGAL4UAS promoter, a C-terminal 3xFLAG epitope (AddGene #50308) and a Nos-terminator (AddGene #50266) into an L1 vector (AddGene #48000). This inducible ETTIN-FLAG cassette was combined with a dexamethasone inducer cassette (containing CD-GVG under a 35S promoter) and a phosphinothricin-resistance cassette in an L2 binary vector (AddGene #48015). The construct was transformed into Agrobacterium tumefaciens strain GV3101 by electroporation, followed by floral dip-mediated plant transformation into Arabidopsis Col-0. For the DARTS assay, seeds were surface sterilised using chlorine gas and germinated in 25 mL liquid ½ MS medium containing 1% sucrose in 100 mL Erlenmeyer flasks under gently shaking. ETTIN-FLAG expression was induced 7 days post germination by adding 10 µM dexamethasone to the growth medium and seedlings were harvested 48 hr post induction. Total plant material was ground in liquid nitrogen and resuspended in protein extraction buffer (25 mM Tris-HCl, pH 7.5; 150 mM NaCl; 0.1% IGEPAL CA-630 and Roche cOmplete protease inhibitor cocktail, EDTA-free) in a 1:1 (w/v) ratio, followed by a centrifuging step to discard the plant debris. Protein concentration was determined with Quick Start Bradford 1x reagent (Bio-Rad) and adjusted to 5 mg/mL by adding extraction buffer. The lysate was then split into different reaction tubes and incubated with the respective chemical (DMSO mock, IAA or BA) at the indicated concentrations for 30 min at room temperature with slow mixing. The treated aliquots were further aliquoted and mixed with different dilutions of Pronase (Roche), prepared in Pronase buffer (25 mM Tris-HCl, pH 7.5 and 150 mM NaCl) to achieve the aimed for ratio of total enzyme to total protein. After incubation for 30 min at room temperature, the proteolytic digestion was terminated by adding protease inhibitor cocktail (cOmplete, Roche) and the samples were kept on ice for 10 min. The protein samples were then mixed with 4x NuPAGE LDS sample buffer (Invitrogen) and heated at 80°C for 10 min. The protein samples were analysed by Western blot (TGX gels Bio-Rad, TurboTransfer Bio-Rad PVDF membranes), visualized using an anti-FLAG-HRP antibody (1:1000 – Sigma A8592). HRP activity was detected using the Supersignal Western Detection Reagents (Thermo Scientific) and imaged with a GE Healthcare Amersham 600 RGB system.

Protein production

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The ES domain, ES388-594, protein was isotopically labelled in preparation for NMR analysis. The ES domain was expressed for as a fusion protein with a 6x Histidine tag in minimal media with 15N ammonium chloride. The 15N isotope labelling of the expressed protein involved a 125-fold dilution of cell culture in enriched growth media into minimal media with 15N ammonium chloride and grown for 16 hr (37 °C / 200 rpm); followed by a further 40-fold dilution into minimal media for the final period of cell growth and protein expression (induced with L-arabinose 0.2 % w/v / 18 °C / 200 rpm and grown for a further 12 hr). The fusion protein was isolated from soluble cell lysate by Co-NTA affinity chromatography with two His-Trap 1 mL TALON Crude columns (GE Healthcare Life Sciences, 28953766). Chromatography buffers contained sodium phosphate 20 mM pH 8.0, NaCl 500 mM and either no-imidazole or 500 mM imidazole for wash and elution buffers respectively. The majority of the non-specifically bound protein was removed by passing 20 mL of the wash buffer through the columns. The protein eluted on a gradient of increasing imidazole concentration of up to 30% elution buffer over 20 mL.

For ITC, the ES388-594 protein was grown in LB medium, cells harvested and lysed as described above. After centrifugation of the sonicated cell suspension, the pellet was resuspended and washed in Wash Buffer (50 mM Tris-HCl, pH 8.0; 0.1 mM EDTA; 5% (v/v) Glycerol; 0.1 mM DTT; 50 mM NaCl; 2% (w/v) NaDOC) for 1 hour at 4°C followed by centrifugation (11,000 g for 20 min at 4 °C). The resulting pellet was again washed in Wash Buffer and centrifuged as described previously. The pellet was resuspended in 20mL of Solubilisation Buffer (50 mM Tris-HCl, pH 8.0; 0.1 mM EDTA; 5% (v/v) Glycerol; 0.1 mM DTT; 50 mM NaCl; 0.25% Sarkosyl (N-Lauroyl sarcosine) and stirred at 4°C for 1h. Subsequently the resuspension was dialysed overnight in 2L Dialysis Buffer (50 mM Tris-HCl, pH 8.0; 0.1 mM EDTA; 5% (v/v) Glycerol; 0.1 mM DTT; 50 mM NaCl). The Dialysis Buffer was exchanged for fresh Dialysis Buffer after at least 2 hours of dialysis. After dialysis the lysate was centrifuged (11,000 g for 20 min at 4 °C). The resulting supernatant contained the required protein which was concentrated to 2 mL using Amicon Ultra-15 centrifugal filters with a 10kDa cut-off (Millipore (UK) Ltd.) and further purified using size exclusion chromatography. The protein was eluted in Chromatography Buffer and concentrated to 2 mL as previously described. Protein concentration was determined with Quick Start Bradford 1x reagent (Bio-Rad) and adjusted using Chromatography Buffer.

HSQC NMR

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The ES domain, ES388-594, protein was analysed by NMR at 5°C under reducing conditions (DTT 10 mM), buffered at pH 8.0 (Tris 20 mM). 1H-15N HSQC was performed at 950 MHz, TCI probe, Bruker following the parameters described in Figure 1—figure supplement 1. Concentrations used were 50 µM ES388-594 and 500 µM IAA ligand (i.e. 1:10).

Isothermal titration calorimetry (ITC)

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ITC was carried out on a MicroCal PEAQ-ITC (Malvern) at 25°C in a Buffer A (sodium phosphate 20 mM, pH 8.0; NaCl 500 mM). Ligand (2 mM IAA) was injected (19 × 4.0 μl) at 150 s intervals into the stirred (500 rpm) calorimeter cell (volume 270 μl) containing 50 µM ES388-594 protein. Titration of Buffer A into 50 µM ES388-594 protein and IAA (2 mM) into Buffer A served as negative controls. Measurements of the binding affinity of all the titration data were analysed using the MicroCal Software (Malvern).

Accessions

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ETT, AT2G33860; TPL, AT1G15750; TPR1, AT1G80490, TPR2, AT3G16830; TPR3, AT5G27030; TPR4, AT3G15880; HDA6, AT5G63110; HDA19, AT4G38130; HEC1, AT5G67060; PID, AT2G34650; WUS, AT2G17950.

References

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    Auxin and ETTIN in Arabidopsis gynoecium morphogenesis
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    ETTIN patterns the Arabidopsis floral meristem and reproductive organs
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    5. KA Feldmann
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    Arabidopsis gynoecium structure in the wild and in ettin mutants
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Decision letter

  1. Jürgen Kleine-Vehn
    Reviewing Editor; University of Natural Resources and Life Sciences, Austria
  2. Christian S Hardtke
    Senior Editor; University of Lausanne, Switzerland

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This is an outstanding study that deepens our mechanistic understanding of a non-canonical auxin perception mechanism and we are very pleased that eLife can provide a forum for this exciting set of data.

Decision letter after peer review:

Thank you for submitting your article "Direct ETTIN-auxin interaction controls chromatin state in gynoecium development" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Christian Hardtke as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

While both reviewers highlighted the suitability of your work for eLife in principle, they also see certain shortcomings, which prevents publication in its present form.

Essential revisions include:

1) The reviewers see a big gap between the in vitro ETT/TPL interaction and the H3K27ac changes in vivo that needs to be bridged. TPL is not the only co-repressor complex that may recruit HDA19 to ETT and ETT interacts with other transcription factors that in turn can also recruit co-repressors. It is currently unclear whether auxin-mediated disruption of the ETT/TPL interaction directly induces changes in HDA19 interaction or chromatin recruitment. This should be tested directly in the presence of auxin in transient interaction assays and by ETT/TPL/HDA19 ChIP +/- IAA at target loci (see further comments to this aspect by reviewer #1).

2) The reviewers also request additional experiments, consolidating the auxin binding to ETT. They encourage the authors to carry out additional MST analyses with ideally using full length ETTIN vs ettin mutant protein (W505). Please quantify binding ETT binding to auxin (see further comments to this aspect by reviewer #2).

Please, see also additional comments by the reviewers below, which may guide you to further strengthen your manuscript.

Reviewer #1:

This study picks up where Simonini et al., 2016 had left off, which had showed that multiple TFs including IND interact with ETT in an auxin -dependent manner. ETT-IND were found to repress PID expression based on in situ hybridization in mutant gynoecia. Mutations in IND that disrupted interaction with ETT led to reduced accumulation of PID, which was interpreted as a role in transcriptional activation of PID by IND and ETT in the absence of auxin.

As I am not an expert on HSQC-NMR or ITC, I will let other reviewers comment on these data. The authors show that nicely TIR1 is neither necessary nor sufficient for HEC1 and PID are induction, suggesting that Aux/IAA degradation is not required for this processes. Critiques that should be addressed are in the numbered/lettered paragraphs below.

1) They next test physical interaction between ETT and TPL/TPRs in Y2H and show it is abolished by addition of IAA. They also test interaction by co-IP in 35S:OE lines after transient transfection into N. benthamiana. In plants, the IAA treatment seems to lead to a both strong reduction in TPL accumulation as well as interaction. This needs to be addressed. It is moreover critical to assess the HDA19 interaction with ETT in N. benthamiana. Using ETT and HDA19 co-transfection into plant cells, ETT was recently shown to interact with HDA19 via endogenous co-repressors (such as TPL or SEU) by co-IP and by BiFC (Chung et al., 2019). In addition, the ETT interaction partner FIL also interacts with HDA19 (likely via different co-repressors). So it is critical to assess whether auxin disrupts ETT complex formation with HDA19 to show that the auxin disrupted ETT/TPL interaction is important for HDA19 association with ETT. As an aside, it should be mentioned in the text that ETT/FIL were shown to recruit HDA19 to STM in the above-mentioned study (Chung et al., 2019).

In ett or tpl /tpr mutant gynoecioa, auxin was not able to induce HEC1 and PID, this was true to a lesser degree for HEC1 in the hda19 mutant. In addition, HEC1 and PID expression is elevated in ett, tpl and hda19 mutants relative to wt. Although not based on inducible mutants, these data suggest that in the wild type ETT and TPL/R repress HEC1 and IND.

2) Figure 5 attempts to show a possible mechanisms for the observed effects by using chromatin immunoprecipitation of ETT, HEC1 and HDA19. There are several issues with the data provided.

a) First, from a technical perspective, the authors should show the factor binding as% input (to give the reader an idea of the occupancy compared to other TFs or chromatin regulators). More importantly, H3K27ac should be presented as fraction of H3. The latter is critical as a change in histone modification is only informative if we know that the nucleosome occupancy has not changed.

b) Second, there was no test for a possible change in TFL or HDA19 occupancy upon auxin treatment at PID or HEC1, although this is a key tenet of their model. ETT occupancy at PID and HEC1 should also be assayed plus minus auxin.

Third, the authors reported lower H3K27ac in the non auxin-treated wild type. This fits with the reduced gene expression, yet cause and consequence are not clear – especially in the absence of test of disruption of ETT- HDA19 interaction by auxin or HDA19 occupancy at the loci being tested. This is important, because changes in gene expression lead to changes in histone modifications and vice -versa.

3) Finally, the title and main conclusion are misleading and oversell the findings.

a) According to the author's data and with some additional support as specified above, the manuscript could show that ETT serves to recruit TPL and HDA19 in the absence of IAA. In the presence of IAA this complex may fall apart. This does not show that ETT toggles between activation and repression. It would show that ETT 's ability to recruit co-repressors is abrogated by auxin.

b) I know the authors suggested this activator/repressor model in the Simonini paper based on dominant mutant proteins, which are tricky to interpret. Besides there being no evidence for such a dual role of ETT in the current paper, I think it is important to remember that the ett loss of function mutant has elevated PID and HEC1. This suggest ETT acts as a repressor. Likewise, for genes bound by MP where loss of MP function results in loss-of-expression, MP is an activator, even though MP does associate with different types of chromatin regulators in the absence or presence of auxin at these loci.

Reviewer #2:

The authors deal with the idea that ETT is a particular ARF protein, that engages directly not only other transcriptional regulators, but the family of TPL co-repressors, and more importantly binds directly auxin. Two previous manuscripts from the same research group (Simonini et al., 2016 and Simonini et al., 2017) had brought up the very intriguing hypothesis that ETT-binds auxin by a non-canonical mechanism, independently of TIR1/AFB-AUX/IAA auxin perception. The hypothesis is fascinating, and the authors previously used genetics, plant physiology and developmental genetics to validate their claim.

In Kuhn et al., the authors seek to: show biochemically direct auxin binding by ETT; connect the ETT "auxin binding role" to specific non-canonical gene expression; bring evidence of ETT directly binding TPL and TPRs for auxin-triggered responses in the gynoecium; show that ETT interactions with TPL and histone deacetylases are sensitive to auxin levels, regulating thereby HEC1 and PID expression. Kuhn et al. work is good and straightforward, in the sense that their hypotheses were addressed by few, but robust experiments. The manuscript would tremendously profit from additional biochemical or biophysical experiments though, using preferably the full length of ETT, to evidence auxin binding. Is that feasible?

Since the authors identified W505 as important putative auxin binding site, why was not an ETT W505 mutant version, and other mutations, tested in iTC or MST, for instance? This reviewer is concerned that since the interaction between ETT and auxin is weak, one could almost account for random peptide interaction, via charges or the similar, for the poor ITC data. Can the authors please quantify binding?

– Along the same lines and about Figure 1 and interactions experiments: Why did the authors not estimate the binding affinity or Kd of ETT-IAA? could the weak (not saturable) ETT-IAA signal be the result of transient peptide interactions?

– When the authors implemented the synthetic auxin-TIR1 tool developed in Keiko Torii's lab, they showed that ETT interaction is highly specific for IAA, but not cvxIAA even though the interaction occurs in an IDR. Can the authors please explain the findings?

– Have the authors evaluated whether there is any sequence conservation for IAA binding around ETT W505?

– Can the authors make the binding comparison of ETT-IAA vs. ETT-BA, or vs. ETT-other auxins, or ETT-tryptophan via iTC in order to make a more robust claim about auxin binding?

– Regarding Model Figure 5D. Do the authors have any hypothesis about what happens with ETT upon auxin binding?

Specific comments:

– The Abstract includes references, is that not a bit unusual?

– Abstract fifth sentence: Sentence too long. Split and/or rephrase

– Abstract final sentence: Giving the outcome of the paper, I recommend to state "ETT might be able to switch chromatin locally…”

– Introduction first paragraph: The word "ubiquitinylation" is incorrect, as it does not refer to ubiquitin conjugation of proteins. Please use "ubiquitination" or "ubiquitylation" instead

– Please rephrase the statement about ES domain being disordered, as it is disconnected, and the reason for bringing it up on that section of the Introduction is not clear.

– The authors used the word "sensitivity", which is a broad claim, given they used a single IAA concentration of 100 µM in their yeast-two hybrid assay.

– HSQC is not the typical experimental approach to proof an X ligand binds to a putative receptor. If this approach is implemented because of the solely disordered nature of ETT, please say it so or rephrase.

– "Helical character" indicated by what? It would be nice to have a visualization help for non-NMR experts.

– “…found that a number of residues shifted…". Please elaborate.

– Why did the authors use benzoic acid (BA) as a control ligand in their experiments? Why not to use tryptophan as the obvious non-active auxin precursor?

– "The HSQC experiment demonstrates that ETT binds IAA directly". I recommend the authors to be cautious here with this statement. While high-field NMR is a powerful tool for investigating transiently forming protein-ligand complexes and for the identification of protein binding partners, there are a number of strategies that the authors must implement first that will specifically enable the physicochemical characterization of ETT-IAA interactions.

– iTC experiments are unfortunately weak e.g. no saturation reached, extremely low signal to noise ratio. Therefore, the authors again should be careful about their statements on direct biochemically relevant ETT-IAA binding.

– Please specify if the ccvTIR1 can accommodate the native IAA.

– Please use here "…independently of the canonical TIR1/AFB1-5 signalling pathway."

– Why IAA = auxin? This is unnecessary and not accurate, stick to "IAA" please.

– Could the authors elaborate on a hypothesis for the role of TPR4? Could its action and interaction with ETT or an ETT-like protein in a different tissue be also auxin sensitive?

– "low levels of auxin" do not maintain per se ETT-TPL/TPR2 associations. Please rephrase.

– The authors have referred twice about the "instantly reversible" event. What do they mean by this? There is no data on timing! "Instantly" as when a [IAA] threshold has been reached?

– The inclusion of an auxin insensitive ett mutant might have shed light on the true nature of the ETT-IAA responsive genes.

Comments on Figures:

– Figure 1. Which IAA concentration was used for these experiments? Please include relevant info in the Figure legend.

– Figure 1. The color of NMR data is hard to distinguish, particularly the blues and the blacks.

Figure 1D-F. Are the control panels E and F necessary as extra figure? Cannot they be included in the D?

– Figure 2. Colors in panel d are difficult to be distinguished. Please recolor.

– Figure 3A. Please improve, resize and label residues. Mark and depict W505.

– Figure 3B. What is WTL? Give info in Legend, please.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "Direct ETTIN-auxin interaction controls chromatin states in gynoecium development" for consideration by eLife. Your article has been reviewed by one peer reviewers, and the evaluation has been overseen by Reviewing Editor Jürgen Kleine-Vehn and Christian Hardtke as the Senior Editor The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our policy on revisions we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.

Both reviewers agreed that you convincingly revised your manuscript and that your manuscript is in principle ready for publication. Reviewer #1 asks you to modify some of your statements and your overall model depiction (see point 2 below).

Reviewer #1:

The authors have addressed the majority of my concerns. I do not recommend additional experiments, but provide a clarification based on the author response. In addition, I ask for changes in the text and model to address my comments in point 2 below.

Previous review

1) It is moreover critical to assess the HDA19 interaction with ETT in N. benthamiana. Using ETT and HDA19 co-transfection into plant cells, ETT was recently shown to interact with HDA19 via endogenous co-repressors (such as TPL or SEU) by co-IP and by BiFC (Chung et al., 2019). In addition, the ETT interaction partner FIL also interacts with HDA19 (likely via different co-repressors). So it is critical to assess whether auxin disrupts ETT complex formation with HDA19 to show that the auxin disrupted ETT/TPL interaction is important for HDA19 association with ETT.

Authors: We are confused about the first sentence of this section that HDA19 interaction with ETT in N. benthamiana must be assessed. As mentioned, this experiment was recently published in Chung et al., 2019, and showed that HDA19 and ETT do not interact directly, but via other proteins. Unless we have misunderstood the reviewer's comment, we do not see the need to carry this out again.

Reviewer: N. benthamiana assays also probe indirect interactions between proteins via host proteins. Perusal of the pulldowns in Chung et al. does indeed reveal a weak HDA19 pulldown by ETT (and a very strong ARF4 pulldown by HDA19). BiFC using Arabidopsis protoplasts showed strong (indirect) interactions between ETT and HDA19. The current study suggests that these indirect interactions should be impaired in the presence of high levels of auxin.

Current review

2) The authors should reconcile their data with the previously published data that implicates ETT/ARF4/FIL in recruitment of HDA19 containing co-repressor complexes to repress KNOX genes in regions of local auxin maxima at the inflorescence shoot apex. In looking at the literature and the current manuscript, a difference in effective auxin levels became apparent. The authors used a rather high auxin concentration to disrupt the ETT – HDA19 co-repressor complex in planta: 100 µM IAA for Figure 4 and Figure 5. By contrast 10 µM IAA is commonly used in inflorescences. Gynoecia may indeed have higher auxin levels.

This suggests a different model and a different conclusion from that currently proposed by the authors are needed. At intermediate auxin levels, for example those of the inflorescence shoot apex, ETT (together with other TFs) is able to recruit HDA19-containing co-repressors, while at higher auxin levels, those present in the gynoecium, this interaction is blocked. The authors should revise the Discussion and their model accordingly.

https://doi.org/10.7554/eLife.51787.sa1

Author response

Essential revisions include:

1) The reviewers see a big gap between the in vitro ETT/TPL interaction and the H3K27ac changes in vivo that needs to be bridged. TPL is not the only co-repressor complex that may recruit HDA19 to ETT and ETT interacts with other transcription factors that in turn can also recruit co-repressors. It is currently unclear whether auxin-mediated disruption of the ETT/TPL interaction directly induces changes in HDA19 interaction or chromatin recruitment. This should be tested directly in the presence of auxin in transient interaction assays and by ETT/TPL/HDA19 ChIP +/- IAA at target loci (see further comments to this aspect by reviewer #1).

We agree that this direct connection between ETT/TPL and HDA19/H3K27ac was not established in the original submission. We performed the ChIP experiments suggested showing that the interaction of both HDA19 and TPL with the promoters of HEC1 and PID is disrupted in the presence of IAA, while ETT binding remains (as previously shown – Simonini et al., 2016). These data support that HDA19 is recruited by ETT via TPL and agree with the disruption of protein interactions between ETT and TPL by IAA. Whilst HDA19 does not directly interact with ETT (Chung et al., 2019), the data support that HDA19 forms part of the repressive ETT/TPL complex that is disrupted by IAA. The data are presented in the revised manuscript.

We did not carry out the ETT-HDA19 interaction assay in N. benthamiana but refer to Chung et al., 2019, where our experiments on this are published. Since no interaction was observed, we suggest it may not be that informative to carry this out +/- IAA.

2) The reviewers also request additional experiments, consolidating the auxin binding to ETT. They encourage the authors to carry out additional MST analyses with ideally using full length ETTIN vs ettin mutant protein (W505). Please quantify binding ETT binding to auxin (see further comments to this aspect by reviewer #2).

Over several years, we tried a range of expression systems and hosts to produce full-length ETT for biochemical studies with no success. Additional attempts could take a long time with no guarantee for success. However, we have previously shown that the ES domain is sufficient for the IAA-sensitivity (Simonini et al., 2016; Simonini et al., 2018) in agreement with numerous other studies reporting a biophysical interaction using an individual protein domain in isolation (e.g. Martin-Arevalillo et al. 2017 PNAS 114, 8107-12; Wild et al. (2016) Science 352, 986-90).

We have not been able to improve on the ITC data presented in the first version. This is due to different factors. Firstly, even with a further optimisation of ES-domain production, we are unable to obtain large enough amounts of fully soluble and stable protein. Secondly, the protein we use seems to aggregate during the ITC measurements making it difficult to obtain a Kd value. This is now discussed in the revised manuscript.

In the revised manuscript, we instead include an additional independent experiment using the DARTS assays providing additional evidence for the direct interaction between ETT and IAA (presented in Figure 1A).

Finally, we also show that the tryptophan in position 505 (W505) is required for the IAAsensitivity of ETT in both Y2H and transient assays in N. benthamiana.

Reviewer #1:

1) They next test physical interaction between ETT and TPL/TPRs in Y2H and show it is abolished by addition of IAA. They also test interaction by co-IP in 35S:OE lines after transient transfection into N. benthamiana. In plants, the IAA treatment seems to lead to a both strong reduction in TPL accumulation as well as interaction. This needs to be addressed.

We agree with the reviewer that the previous figure had lower levels of TPL at higher concentrations of IAA. We have carried this out several times without an effect on TPL levels, but showing the reduction in ETT-TPL interaction. We have now included the results from a more representative experiments (Figure 3C,D, Figure 3—figure supplement 3).

It is moreover critical to assess the HDA19 interaction with ETT in N. benthamiana. Using ETT and HDA19 co-transfection into plant cells, ETT was recently shown to interact with HDA19 via endogenous co-repressors (such as TPL or SEU) by co-IP and by BiFC (Chung et al., 2019). In addition, the ETT interaction partner FIL also interacts with HDA19 (likely via different co-repressors). So it is critical to assess whether auxin disrupts ETT complex formation with HDA19 to show that the auxin disrupted ETT/TPL interaction is important for HDA19 association with ETT. As an aside, it should be mentioned in the text that ETT/FIL were shown to recruit HDA19 to STM in the above-mentioned study (Chung et al., 2019).

As mentioned above, to assess whether auxin disrupts ETT complex formation with HDA19, we have performed ChIP with HDA19/TPL/ETT +/-IAA and found that the interaction of HDA19 and TPL with the HEC1 and PID promoters is sensitive to IAA, while ETT remains bound independently of IAA levels. This supports the model that ETT recruits HDA19 via interactions with TPL and that the ETT/HDA19/TPL complex is disrupted by IAA. These data are included in the manuscript (Figure 5A-C).

We are confused about the first sentence of this section that HDA19 interaction with ETT in N. benthamiana must be assessed. As mentioned, this experiment was recently published in Chung et al., 2019, and showed that HDA19 and ETT do not interact directly, but via other proteins. Unless we have misunderstood the reviewer’s comment, we do not see the need to carry this out again.

We agree with the reviewer about specifically mentioning the previous finding that ETT is required to recruit HDA19 to the STM promoter as reported in Chung et al., 2019. This is now included in the revised version of the manuscript.

In ett or tpl /tpr mutant gynoecioa, auxin was not able to induce HEC1 and PID, this was true to a lesser degree for HEC1 in the hda19 mutant. In addition, HEC1 and PID expression is elevated in ett, tpl and hda19 mutants relative to wt. Although not based on inducible mutants, these data suggest that in the wild type ETT and TPL/R repress HEC1 and IND.

2) Figure 5 attempts to show a possible mechanisms for the observed effects by using chromatin immunoprecipitation of ETT, HEC1 and HDA19. There are several issues with the data provided.

a) First, from a technical perspective, the authors should show the factor binding as% input (to give the reader an idea of the occupancy compared to other TFs or chromatin regulators). More importantly, H3K27ac should be presented as fraction of H3. The latter is critical as a change in histone modification is only informative if we know that the nucleosome occupancy has not changed.

We agree with the technical concerns raised by the reviewer and have revised the figure as suggested (Figure 5D, E).

b) Second, there was no test for a possible change in TFL or HDA19 occupancy upon auxin treatment at PID or HEC1, although this is a key tenet of their model. ETT occupancy at PID and HEC1 should also be assayed plus minus auxin.

We agree with this point and have performed additional ChIP experiments plus/minus auxin to test ETT, TPL and HDA19 occupancy at PID and HEC1. The new data have been included in the revised figure (Figure 5A-C).

Third, the authors reported lower H3K27ac in the non auxin-treated wild type. This fits with the reduced gene expression, yet cause and consequence are not clear – especially in the absence of test of disruption of ETT- HDA19 interaction by auxin or HDA19 occupancy at the loci being tested. This is important, because changes in gene expression lead to changes in histone modifications and vice -versa.

We agree that this is an important point and have addressed experimentally the HDA19 occupancy at PID and HEC1 by ChIP as requested by the reviewer above. Moreover, we have inserted a sentence to highlight that no direct interaction between ETT and HDA19 was detected in both Y2H and Co-IP (Chung et al., 2019); however, an indirect interaction via a co-repressor was suggested.

3) Finally, the title and main conclusion are misleading and oversell the findings.

a) According to the author's data and with some additional support as specified above, the manuscript could show that ETT serves to recruit TPL and HDA19 in the absence of IAA. In the presence of IAA this complex may fall apart. This does not show that ETT toggles between activation and repression. It would show that ETT 's ability to recruit co-repressors is abrogated by auxin.

We do not agree that the title is misleading and would prefer to keep as is. However, we have modified the concluding remarks to take the reviewer’s comments on repressive/de-repressive ability of ETT on board. This has led to a section in which we suggest the possibility that auxin either renders ETT a passive DNA-binding protein or has a role in recruiting activating components.

b) I know the authors suggested this activator/repressor model in the Simonini paper based on dominant mutant proteins, which are tricky to interpret. Besides there being no evidence for such a dual role of ETT in the current paper, I think it is important to remember that the ett loss of function mutant has elevated PID and HEC1. This suggest ETT acts as a repressor. Likewise, for genes bound by MP where loss of MP function results in loss-of-expression, MP is an activator, even though MP does associate with different types of chromatin regulators in the absence or presence of auxin at these loci.

As above to 3a.

Reviewer #2:

[…] In Kuhn et al., the authors seek to: show biochemically direct auxin binding by ETT; connect the ETT "auxin binding role" to specific non-canonical gene expression; bring evidence of ETT directly binding TPL and TPRs for auxin-triggered responses in the gynoecium; show that ETT interactions with TPL and histone deacetylases are sensitive to auxin levels, regulating thereby HEC1 and PID expression. Kuhn et al. work is good and straightforward, in the sense that their hypotheses were addressed by few, but robust experiments. The manuscript would tremendously profit from additional biochemical or biophysical experiments though, using preferably the full length of ETT, to evidence auxin binding. Is that feasible?

Despite using a large range of different expression systems over several years, we are unable to produce full length ETT in any recombinant system. As described in Simonini et al., 2018, we can make an almost full version of the Cterminal ES domain (only lacking 14 amino acids at the terminus), which is sufficient for the ETT-sensitivity.

Since the authors identified W505 as important putative auxin binding site, why was not an ETT W505 mutant version, and other mutations, tested in iTC or MST, for instance? This reviewer is concerned that since the interaction between ETT and auxin is weak, one could almost account for random peptide interaction, via charges or the similar, for the poor ITC data. Can the authors please quantify binding?

As explained above, we have not been able to improve on our ITC data despite several attempts (including developing a fusion protein that stayed in solution, but had lost ability to bind IAA in this assay). However, we agree with the reviewer on exploiting the potential importance of the tryptophan in position 505 (W505) in the ETT-auxin interaction. Therefore, we developed mutant versions substituting tryptophan for alanine (W505A) and tested them in interaction assays ± IAA (Y2H and Co-IP with TPL). In agreement with a central role of W505, the ETT(W505A)-TPL interaction was not disrupted by IAA (Figure 3B, D).

– Along the same lines and about Figure 1 and interactions experiments: Why did the authors not estimate the binding affinity or Kd of ETT-IAA? could the weak (not saturable) ETT-IAA signal be the result of transient peptide interactions?

Possibly due to insufficiently high protein concentration and/or aggregation, we were unable to obtain ITC data allowing quantification of the binding affinity.

– When the authors implemented the synthetic auxin-TIR1 tool developed in Keiko Torii's lab, they showed that ETT interaction is highly specific for IAA, but not cvxIAA even though the interaction occurs in an IDR. Can the authors please explain the findings?

Without a protein structure it is hard to explain the specific requirement for IAA over other auxinic compounds in this pathway. However, this is not just IAA vs cvxIAA. We have previously reported that neither NAA nor 2,4-D are able to mediate the effect on ETTprotein interactions (this paper and Simonini et al., 2016).

– Have the authors evaluated whether there is any sequence conservation for IAA binding around ETT W505?

This is an excellent point. We have found that the tryptophan in position 505 is highly conserved among the 25 ETT proteins that we have identified. In contrast, very little sequence conservation is observed among other residues in that region. We have included a sequence logo of the W505 region in the alignment in Figure 3—figure supplement 1A to point this out.

– Can the authors make the binding comparison of ETT-IAA vs. ETT-BA, or vs. ETT-other auxins, or ETT-tryptophan via iTC in order to make a more robust claim about auxin binding?

In the revised manuscript, we use IAA vs BA in the DARTS and NMR experiments and IAA vs. NAA in the Y2H and Co-IP in N. benthamiana. We did not test all variations in ITC due to difficulty in obtaining sufficient protein as described above.

– Regarding Model Figure 5D. Do the authors have any hypothesis about what happens with ETT upon auxin binding?

Structurally, we do not yet have any idea what changes are occurring in the ETT protein upon IAA binding, although, our recent data included in this revised manuscript suggest that the tryptophan in position 505 may play an important role. Moreover, in response to the comments from reviewer 1, we have modified the speculations to include the possibility that ETT functions primarily as a repressor and that IAA binding induces a conformational change in ETT to make it unable to bind the co-repressive complex. An alternative hypothesis is that IAA-bound ETT is required for de-repression by recruiting activators of gene expression. Both possibilities are presented in the revised version.

Specific comments:

– The Abstract includes references, is that not a bit unusual?

We are happy to exclude if the journal requests this. However, it seems some eLife papers have references in the Abstract whereas others do not.

– Abstract fifth sentence: Sentence too long. Split and/or rephrase

We agree and have modified the sentence.

– Abstract final sentence: Giving the outcome of the paper, I recommend to state "ETT might be able to switch chromatin locally…”

We have modified this sentence.

– Introduction first paragraph: The word "ubiquitinylation" is incorrect, as it does not refer to ubiquitin conjugation of proteins. Please use "ubiquitination" or "ubiquitylation" instead

This has been corrected.

– Please rephrase the statement about ES domain being disordered, as it is disconnected, and the reason for bringing it up on that section of the Introduction is not clear.

We agree and have modified the sentence.

– The authors used the word "sensitivity", which is a broad claim, given they used a single IAA concentration of 100 µM in their yeast-two hybrid assay.

We disagree and prefer to use the term ‘sensitivity’. We have previously shown that the interactions of ETT with the IND TF occurs with several IAA concentrations (Simonini et al., 2016).

– HSQC is not the typical experimental approach to proof an X ligand binds to a putative receptor. If this approach is implemented because of the solely disordered nature of ETT, please say it so or rephrase.

NMR was used to investigate the binding of auxin to ETT principally because it is a powerful technique for analysing interactions that are relatively low affinity, as was suspected in this case. Such HSQC experiments examine the interaction from the perspective of the receptor rather than the ligand (as is the case in WaterLOGSY NMR experiments) and provide information on the number of residues of the receptor that are affected by the binding of the ligand, indicated by changes in the position of HN cross peaks within the spectra.

– "Helical character" indicated by what? It would be nice to have a visualization help for non-NMR experts.

The helical character is typically indicated by the dispersed HN cross peaks between 7.8 to 7.0 ppm on the 1H chemical shift axis of the HSQC spectrum. This has now been indicated in Figure 1B, C.

– “…found that a number of residues shifted…". Please elaborate.

Perturbations in the position of the HN cross peaks in the HSQC is due to changes in the local chemical environment of the amide bonds within the protein. This is caused either by being in close proximity to bound ligand and/or due to conformational change to the protein induced by the binding event. These IAA specific induced changes are shown in Figure 1D.

– Why did the authors use benzoic acid (BA) as a control ligand in their experiments? Why not to use tryptophan as the obvious non-active auxin precursor?

Benzoic acid was selected as the control because it has no auxinic activity but like IAA, is a weak acid.

– "The HSQC experiment demonstrates that ETT binds IAA directly". I recommend the authors to be cautious here with this statement. While high-field NMR is a powerful tool for investigating transiently forming protein-ligand complexes and for the identification of protein binding partners, there are a number of strategies that the authors must implement first that will specifically enable the physicochemical characterization of ETT-IAA interactions.

The IAA specific induced changes in the HSQC spectrum is the effect of IAA binding directly to ETT, either by direct interaction with the binding interface or due to more distal conformational changes as a result of binding. This protein-ligand complex is further supported by ITC and DARTS assays. These results do not rule out possibility that the bioactive complex involves further binding partners.

– iTC experiments are unfortunately weak e.g. no saturation reached, extremely low signal to noise ratio. Therefore, the authors again should be careful about their statements on direct biochemically relevant ETT-IAA binding.

We agree with the reviewer and have included text to explain the problems we have experienced with the ITC. In addition to the HSQC-NMR data we now also show data from a DARTS experiment that provide additional evidence of direct interaction (Figure 1A).

– Please specify if the ccvTIR1 can accommodate the native IAA.

This is described in Uchida et al., 2018 and their data show that ccvTIR1 does not respond to IAA. This is now specifically mentioned in the manuscript.

– Please use here "…independently of the canonical TIR1/AFB1-5 signalling pathway."

This has been modified.

– Why IAA = auxin? This is unnecessary and not accurate, stick to "IAA" please.

We have changed to (primarily) use IAA and have specified this earlier in manuscript.

– Could the authors elaborate on a hypothesis for the role of TPR4? Could its action and interaction with ETT or an ETT-like protein in a different tissue be also auxin sensitive?

This is an entirely possible scenario and we have included a sentence in the revised manuscript to that extent.

– "low levels of auxin" do not maintain per se ETT-TPL/TPR2 associations. Please rephrase.

We agree and have modified this sentence.

– The authors have referred twice about the "instantly reversible" event. What do they mean by this? There is no data on timing! "Instantly" as when a [IAA] threshold has been reached?

We have modified the text at both places in response to the reviewer’s comment. In fact, we like the ‘threshold’ formulation so much that we have included this in the revised manuscript.

– The inclusion of an auxin insensitive ett mutant might have shed light on the true nature of the ETT-IAA responsive genes.

We are not entirely sure what the reviewer means by “the true nature of the

ETT-IAA responsive genes”; however, we refer to the paper by Simonini et al., 2016, where expressing a version of the TF IND (INDD30G) that makes the ETT-IND interaction insensitive leads to constitutive repression of PID (shown by in situ hybridisation in the gynoecium).

Comments on Figures:

– Figure 1. Which IAA concentration was used for these experiments? Please include relevant info in the Figure legend.

ES388-594:IAA / 1:10 / 50 µM:500 µM. This has now been indicated in the Figure legend.

– Figure 1. The color of NMR data is hard to distinguish, particularly the blues and the blacks.

Colour of the NMR data has been changed to emphasise the contrast with black ES388-594 only control sample.

Figure 1D-F. Are the control panels E and F necessary as extra figure? Cannot they be included in the D?

We prefer to include the ITC controls as they are to clearly show the difference between them and the result with both ES and IAA.

– Figure 2. Colors in panel d are difficult to be distinguished. Please recolor.

This has been modified.

– Figure 3A. Please improve, resize and label residues. Mark and depict W505.

This has been modified.

– Figure 3B. What is WTL? Give info in Legend, please.

We believe the reviewer is referring to the -W-L-A-H, which indicates Yeast Selection medium lacking Tryptophan (W), Leucin (L), Adenine (A) and Histidine (H) to assess interaction. This is now specified in the legend.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Reviewer #1:

The authors have addressed the majority of my concerns. I do not recommend additional experiments, but provide a clarification based on the author response. […]

Reviewer: N. benthamiana assays also probe indirect interactions between proteins via host proteins. Perusal of the pulldowns in Chung et al. does indeed reveal a weak HDA19 pulldown by ETT (and a very strong ARF4 pulldown by HDA19). BiFC using Arabidopsis protoplasts showed strong (indirect) interactions between ETT and HDA19. The current study suggests that these indirect interactions should be impaired in the presence of high levels of auxin.

We thank the reviewer for clarifying this comment. We believe the new ChIP data provided using a p35S:HDA19:GFP line in presence of auxin also addresses this point showing that auxin leads to dissociation of HDA19 from ETT-target loci, while ETT remains bound.

Current review

2) The authors should reconcile their data with the previously published data that implicates ETT/ARF4/FIL in recruitment of HDA19 containing co-repressor complexes to repress KNOX genes in regions of local auxin maxima at the inflorescence shoot apex. In looking at the literature and the current manuscript, a difference in effective auxin levels became apparent. The authors used a rather high auxin concentration to disrupt the ETT – HDA19 co-repressor complex in planta: 100 µM IAA for Figure 4 and Figure 5. By contrast 10 µM IAA is commonly used in inflorescences. Gynoecia may indeed have higher auxin levels.

This suggests a different model and a different conclusion from that currently proposed by the authors are needed. At intermediate auxin levels, for example those of the inflorescence shoot apex, ETT (together with other TFs) is able to recruit HDA19-containing co-repressors, while at higher auxin levels, those present in the gynoecium, this interaction is blocked. The authors should revise the Discussion and their model accordingly.

This is an excellent point and we agree with the reviewer that different auxin concentrations may have different effects in diverse developmental contexts. It is, moreover, possible that different ETT-containing complexes require different auxin concentration thresholds to generate auxin responses. We have modified the Discussion accordingly (first paragraph); however, we think that Figure 5F depicts our model appropriately.

https://doi.org/10.7554/eLife.51787.sa2

Article and author information

Author details

  1. André Kuhn

    Department of Crop Genetics, John Innes Centre, Norwich Research Park, Norwich, United Kingdom
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2144-8413
  2. Sigurd Ramans Harborough

    Centre for Plant Sciences, Faculty of Biological Sciences, University of Leeds, Leeds, United Kingdom
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0361-5647
  3. Heather M McLaughlin

    Department of Crop Genetics, John Innes Centre, Norwich Research Park, Norwich, United Kingdom
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3020-7964
  4. Bhavani Natarajan

    Department of Crop Genetics, John Innes Centre, Norwich Research Park, Norwich, United Kingdom
    Contribution
    Data curation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6852-7130
  5. Inge Verstraeten

    Institute of Science and Technology, Klosterneuburg, Austria
    Contribution
    Data curation, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7241-2328
  6. Jiří Friml

    Institute of Science and Technology, Klosterneuburg, Austria
    Contribution
    Formal analysis, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8302-7596
  7. Stefan Kepinski

    Centre for Plant Sciences, Faculty of Biological Sciences, University of Leeds, Leeds, United Kingdom
    Contribution
    Formal analysis, Funding acquisition
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9819-5034
  8. Lars Østergaard

    Department of Crop Genetics, John Innes Centre, Norwich Research Park, Norwich, United Kingdom
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Project administration
    For correspondence
    lars.ostergaard@jic.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8497-7657

Funding

Biotechnology and Biological Sciences Research Council (BB/S002901/1)

  • Lars Østergaard

Biotechnology and Biological Sciences Research Council (BB/L010623/1)

  • Stefan Kepinski

Biotechnology and Biological Sciences Research Council (BB/M011216/1)

  • André Kuhn

Biotechnology and Biological Sciences Research Council (BB/P013511/1)

  • Lars Østergaard

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are grateful to Yuli Ding, Yang Dong, Emilie Knight, Mikhaela Neequaye, Nicola Stacey, Sophia Stavnstrup, Billy Tasker-Brown for critical comments on the manuscript, to Keiko Torii and Shinya Hagihara for the ccvTIR1 line and cvxIAA ligand, to Rebecca Mosher for assistance with the phylogenetic analysis of ETT protein sequences, to Thomas Laux for TPL::TPL-GFP and 35S::HDA19-GFP lines and to Salomé Prat for 35S::TPL-HA construct. Our gratitude also goes to Felix Ramos-León, Kelley Gallagher and Mark Buttner for assistance with protein production, Clare Stevenson for ITC support, the Norwich Research Park Bioimaging for skillful assistance and the NMR facility in the Astbury Centre, Faculty of Biological Sciences for access to the 950 MHz and 600 MHz spectrometers funded by the University of Leeds. We thank Arnout Kalverda for assistance with analysing the NMR data.

Senior Editor

  1. Christian S Hardtke, University of Lausanne, Switzerland

Reviewing Editor

  1. Jürgen Kleine-Vehn, University of Natural Resources and Life Sciences, Austria

Publication history

  1. Received: September 11, 2019
  2. Accepted: April 5, 2020
  3. Accepted Manuscript published: April 8, 2020 (version 1)
  4. Version of Record published: April 17, 2020 (version 2)

Copyright

© 2020, Kuhn et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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