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. Author manuscript; available in PMC: 2022 Oct 18.
Published in final edited form as: J Exp Bot. 2022 Apr 18;73(8):2308–2319. doi: 10.1093/jxb/erac019

Auxin analog-induced Ca2+ signaling is not involved in inhibition of endosomal aggregation

Ren Wang 1,2,*, Ellie Himschoot 1,2,*, Matteo Grenzi 3,*, Jian Chen 1,2, Alaeddine Safi 1,2, Melanie Krebs 4, Karin Schumacher 4, Moritz K Nowack 1,2, Daniёl Van Damme 1,2, Ive De Smet 1,2, Danny Geelen 5, Tom Beeckman 1,2, Jiří Friml 6, Alex Costa 3,7,§, Steffen Vanneste 1,5,8,§
PMCID: PMC7612644  EMSID: EMS141012  PMID: 35085386

Abstract

Much of what we know about the role of auxin in plant development derives from exogenous manipulations of auxin distribution and signaling, using inhibitors, auxins and auxin analogs. In this context, synthetic auxin analogs, such as 1-Naphtalene Acetic Acid (1-NAA), are often favored over the endogenous auxin indole-3-acetic acid (IAA), in part due to their higher stability. While such auxin analogs have proven to be instrumental to reveal the various faces of auxin, they displays in some cases distinct bioactivities compared to IAA. Here, we focused on the effect of auxin analogs on the accumulation of PIN proteins in Brefeldin A (BFA)-sensitive endosomal aggregations (BFA bodies), and the correlation with the ability to elicit Ca2+ responses. For a set of commonly used auxin analogs, we evaluated if auxin-analog induced Ca2+ signaling inhibits PIN accumulation in. We found that not all auxin analogs elicited a Ca2+ response, and that their differential ability to elicit Ca2+ responses correlated partially with their ability to inhibit BFA-body formation. However, in tir1/afb and cngc14, 1-NAA-induced Ca2+ signaling was strongly impaired, while 1-NAA’s still could inhibit PIN accumulation in BFA bodies. This demonstrates that TIR1/AFB-CNGC14-dependent Ca2+ signaling does not inhibit BFA body formation.

Keywords: Arabidopsis, Auxin-analogs Calcium, endosomes, root, signaling, trafficking

Introduction

The natural auxin, indole-3-acetic acid (IAA) is a potent regulator of plant development. Therefore, plants have developed rigorous mechanisms to control IAA activity. In many cases, auxin itself activates a feedback regulation on its own activity, via activation of auxin transport, compartimentalisation and conjugation (Adamowski and Friml, 2015; Casanova-Saez et al., 2021; Rosquete et al., 2012). This, and a low stability in medium, has led to a preferential use of more stable, synthetic auxin to study long-term auxin effects on plant development. Despite obvious drawbacks, the use of auxin analogues has been very successful for the identification of multiple components of auxin signaling and transport via forward genetic screens based on resistance to 2,4-D (Estelle and Sommerville, 1987; Lincoln et al., 1990; Maher and Martindale, 1980) and 1-NAA (Ashton et al., 1979). This illustrates how synthetic auxins have been used to drive auxin research forward.

The auxin analogues can thus activate what is now known as the canonical pathway for auxin-induced transcription. Auxin or a synthetic analog stabilizes interaction between TIR1/AFB F-box proteins and Aux/IAA transcriptional co-repressors (Dharmasiri et al., 2005; Kepinski and Leyser, 2005), to trigger the proteolysis of the latter(Nemhauser, 2018), resulting in derepression of ARF transcription factors (Pierre-Jerome et al., 2016; Roosjen et al., 2018). TIR1/AFB-based auxin sensing has also recently emerged in the context of non-transcriptional auxin signaling, explaining auxin-induced membrane depolarization, apoplast alkalinisation, Ca2+ signaling and root growth inhibition (Dindas et al., 2018; Fendrych et al., 2018; Gallei et al., 2020; Li et al., 2021; Serre et al., 2021).

Arabidopsis contains 6 TIR1/AFB family members with overlapping and specific functions (Prigge et al., 2020). Different auxins and auxin analogs have distinct affinities and binding specificities towards either TIR1-type (TIR1, AFB1,2 & 3) or AFB5-type (AFB4 & 5) AFBs (Lee et al., 2014), and can thus be used to selectively activate a subset of them. A well-known example of this is picloram, that selectively activates AFB4 and AFB5, a biochemical activity that is reflected in the picloram resistance of the corresponding mutants (Prigge et al., 2016). In contrast, most other commonly used auxins and auxin analogs, such as IAA, 1-NAA and 2,4-D are promiscuous activators of all TIR1/AFBs (Calderon Villalobos et al., 2012; Kepinski and Leyser, 2005; Lee et al., 2014). In many cases, 2-NAA is used as inactive 1-NAA analog, despite displaying a weak affinity for TIR1-type AFBs (Lee et al., 2014).

The use of auxin analogs also identified effects of auxin on pavement cell morphogenesis (Xu et al., 2010), lipid composition and distribution (Li et al., 2015; Pan et al., 2009; Platre et al., 2019), cell division during lateral root formation (Huang et al., 2019), suppression of auxin biosynthesis (Wang et al., 2020), stability and polarity of PIN proteins (Abas et al., 2006; Baster et al., 2013; Mazur et al., 2020a; Prat et al., 2018; Sauer et al., 2006), and complex effects on the endomembrane system (Narasimhan et al., 2021; Paciorek et al., 2005; Platre et al., 2019; Robert et al., 2010), apoplast acidification (Li et al., 2021; Lin et al., 2021). In most cases, only the auxin perception mechanism has been identified, TIR1/AFB and/or the receptor-like kinase family TRANSMEMBRANE KINASE1-4 (Cao et al., 2019; Huang et al., 2019; Li et al., 2021; Lin et al., 2021; Platre et al., 2019).

An inhibitory effect of auxin on internalization has been proposed in part based on experiments using the synthetic auxin 1-naphtalene acetic acid (1-NAA) in combination with the fungal toxin Brefeldin A (BFA) (Paciorek et al., 2005; Robert et al., 2010). This inhibitor (BFA) is a commonly used cell biological tool to inhibit secretion. It stabilizes the interaction between ADP-Ribosylation Factor (ARF) small GTPases, and their activators, ARF-Guanine-nucleotide Exchange Factors (ARF-GEF), thereby trapping the complex at the membrane surface (Brumm et al., 2020; Renault et al., 2003), and impairing ARF-mediated membrane trafficking. A side-effect of BFA treatment is the formation of so-called BFA-bodies, large endosomal aggregates that are surrounded by Golgi stacks (Robinson et al., 2008), in which endocytosed, as well as de novo synthesized cargoes are concentrated (Jasik and Schmelzer, 2014; Narasimhan et al., 2021). The synthetic auxin, 1-NAA, has a profound effect on the accumulation of PINs in BFA bodies (Paciorek et al., 2005), an effect that is much weaker when using the endogenous auxin IAA (Paciorek et al., 2005; Paponov et al., 2019; Simon et al., 2013). Side by side comparison of the effects of 1-NAA and IAA on the endomembrane system organization and dynamics has revealed that both auxinic molecules interfere with BFA-induced endosomal aggregation, albeit with different efficiency, and neither of them interfered with the incidence and dynamics of clathrin-mediated endocytic events (Narasimhan et al., 2021). On the other hand, unlike IAA, 1-NAA affects the uptake of the endocytic tracer dye FM4-64, and impacted on the organization and identity of endosomal compartments (Narasimhan et al., 2021). These examples indicate that IAA and 1-NAA have differential activities in the cell. A good understanding of the mechanisms underlying the differential activities of both molecules is essential to interpret the physiological relevance of the responses to either molecule.

Calcium is the universal second messenger in living cells, and has a key roles in many cellular processes, including endomembrane trafficking (Himschoot et al., 2017). However, its ubiquitous activity makes it difficult to isolate any specific function. Indeed, auxin-induced Ca2+ signaling has been known for many years, but its physiological role remains poorly understood (Vanneste and Friml, 2013). In recent years, however, critical progress has been made in unraveling auxin-induced Ca2+ signaling. The reduced IAA-induced Ca2+ fluxes tir1/afb2,3 mutants (Dindas et al., 2018), and induction of Ca2+ signals by via orthogonal activation of an engineered TIR1 variant (Li et al., 2021), identified TIR1/AFB family members as auxin sensors for auxin-induced Ca2+ signaling. Two non-selective cation channel subunits CNGC14 and CNGC2 have been identified as critical components of auxin-induced Ca2+ entry (Chakraborty et al., 2021; Dindas et al., 2018; Shih et al., 2015). CNGC14 activity was proposed to participate in root gravitropism (Shih et al., 2015) and root hair development (Brost et al., 2019; Dindas et al., 2018). CNGC2 is well characterized in defense related Ca2+ signaling, and was recently found to control auxin biosynthesis and signaling, and showing auxin-resistance (Chakraborty et al., 2021). While the interrelationship between these components remains to be resolved, these observations provide 3 independent approaches to specifically interfere with auxin-induced Ca2+ signaling, and assess the impact on cellular processes. Here, we used these genetic tools and auxin analogs to assess the role of auxin-analog induced Ca2+ signaling in regulation of PIN aggregation in BFA bodies.

Materials and Methods

Plant Growth Conditions

Arabidopsis thaliana seeds were sterilized by using bleach gas (8mL concentrated HCl to 150mL bleach) overnight, afterwards the seeds were sown on Petri dishes (12 cm X 12 cm) containing sterile half-strength Murashige and Skoog (0.5 x MS) medium (0.5 x MS salts, 0.8% sucrose, 0.5 g/L 2-(N-morpholino) ethanesulfonic acid, pH 5.7, and 1% w/v agar), and grown under continuous light (21°C, continuous light), after 2 days stratification at 4°C in the dark.

Chemicals

The following hormones/chemicals were used: 1-NAA (catalog Nr N0903; Duchefa Biochemie). Brefeldin A (BFA, catalog Nr B6542-25MG-Sigma Aldrich). 2-NAA (catalog Nr N4002-5G-A-Sigma Aldrich). Quinclorac (catalog Nr 36521-250MG-Sigma Aldrich). 2,4-dichlorophenoxy acetic acid (2,4-D, catalog Nr D7299-100G-Sigma Aldrich). Dicamba (catalog Nr 45430-250MG-Sigma Aldrich) and Picloram (catalog Nr 36774-250MG-R-Sigma Aldrich). α-(phenyl ethyl-2-one)-indole-3-acetic acid (PEO-IAA, catalog Nr TA9H97BAEB94-5MG-Sigma Aldrich). Auxinole (catalog Nr BL3H160B7E0C-50MG-Sigma Aldrich). The natural auxin PhenylAcetic Acid (PAA, catalog Nr P16621-5G-Sigma Aldrich) and the non-specific control Benzoic Acid (BA, catalog Nr 242381-25G-Sigma Aldrich). All hormones/drugs were dissolved in 100% dimethylsulfoxide (DMSO; catalog Nr D4540-500ML-Sigma Aldrich).

Plant Lines Used

The Arabidopsis line used as control in this study was Columbia (Col-0) ecotype. We used the following mutants and transgenic lines were described previously: tir1-1 (Ruegger et al., 1998), afb2-3 (Parry et al., 2009), afb3-4 (Parry et al., 2009), tir1-1afb2-3 (Parry et al., 2009), tir1-1afb3-4 (Parry et al., 2009), tir1-1afb1-3afb3-4 (Parry et al., 2009), tir1-1afb2-3afb3-4 (Parry et al., 2009), cngc14-1 and cngc14-2 (Shih et al., 2015), cngc6-1/9-3, cngc6-1,14-4, cngc9-3,14-4 and cngc6-1,9-3,14-14 (Brost et al., 2019), NES-YC3.60 (Krebs et al., 2012), PM-YC3.60-Lti6b (Krebs et al., 2012), CRT-D4ER (Bonza et al., 2013), 4mt-YC3.60 (Loro et al., 2012). The NES-YC3.60 (Krebs et al., 2012) reporter construct was transformed directly into tir1-1afb2-3afb3-4 (Parry et al., 2009) and cngc14-1 (Shih et al., 2015). Transformants were selected based on uniform expression, and expression levels comparable to the control NES-YC3.60 (Fig. 4A). Analyses of the Ca2+ response was done on T2 generation seedling roots showing strong and uniform expression.

Fig. 4. 1-NAA-induced Ca2+ responses involve TIR1/AFB and CNGC14.

Fig. 4

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(A) Comparison of the raw cpVenus/CFP ratios of the off-spring of the selected NES-YC3.60 x tir1afb2afb3 and NES-YC3.60 x cngc14-1 lines used in this study, relative to the original NES-YC3.60 line (WT). The ratios were quantified and plotted as box-and-whisker plots.

(B-C) The Ca2+ dynamics (NES-YC3.60) in WT, tir1afb2afb3, cngc14-1 in response to a pulse of 10μM 1-NAA using wide-field illumination. Data are presented as means + S.D. n=5 (B). The maximal NES-YC3.60 responses were quantified and plotted as box-and-whisker plots (C).

In (A) and (C) all experimental points are plotted and their distribution represented as a box that extends from the 25th to 75th percentiles. The central line in the boxes are at the respective medians. (Unpaired Student’s t-test; non-significant (ns) P > 0.05, * P<0.05, **P<0.01, *** P<0.001).

(D) Expression pattern of CNGC14 in the meristem. Screenshot from https://bioit3.irc.ugent.be/plant-sc-atlas/root using CNGC14 as input. Corresponding annotation of the tissue-specific cell clusters was implemented as outlined in the figure.

(E) Analysis of the 1-NAA-induced Ca2+ response of NES-YC3.60 in WT and cgnc14-1 via laser scanning confocal microscopy, in the same ROI as taken in the wide field illumination experiments. The focus was set to capture all root tissues with a z-depth of 3.77μm.

(F) Kymograph analysis of NAA-induced Ca2+ signaling over in WT and in cngc14-1 based on the movies that were used to construct the graph in Fig. 4E. Fluorescence ratios across the indicated dotted line of a central confocal section (a to b; 100μm) were plotted over time. A white line in the kymographs indicates the pulse of 10 μM 1-NAA. The color-coded scale indicates NES-YC3.60 fluorescence ratios from low to high.

Immunodetection

The seedlings used for immunodetection are 3-day-old seedlings, pre-treated in liquid 0.5 x MS for 30 minutes, followed by a 1h (co)treatment with BFA (25μM) or BFA (25μM) /NAA (10μM) by transfer to fresh 0.5 x MS with the respective chemicals. The samples were fixed by paraformaldehyde (4%) in PBS for 1 hour in vacuum. The following steps of the immunostaining were performed by the immuno-robot InsituPro Vsi II (Intavis), as described by Sauer et al. (2006). In brief, immediately after fixation, the samples were transferred to the 30-well rack of the robot and subjected to 6 washes (3 in PBS pH 7.4+ 0.1%Triton X-100 (PBS-T); 3 in distilled water + 0.1%Triton X-100; 5 min each), cell wall digestion (1.5% Driselase in PBS, 30min at 37°C;), 3 washes (PBS-T, 5min), permeabilization (3% NP-40, 10% DMSO in PBS; 30min room temperature), 3 washes (PBS-T, 5min), blocking solution (3% BSA in PBS, 1h 37°C), primary antibody solution (primary antibodies diluted in blocking solution, 4h 37°C), 5 washes (PBS-T, 5min), secondary antibody solution (secondary antibodies diluted in blocking solution, 3h, 37°C), 5 washes (PBS-T, 5min) and 5 washes (distilled water, 5 min). The dilutions of the primary antibodies used are: goat anti-PIN1 (1:600) (sc-27163, SantaCruz). The dilutions of the secondary antibodies used are: AlexaFluor488 donkey anti-goat (1:600) (A-11055, ThermoFisher). Seedlings were mounted in AF1 antifadant (Citifluor). A Leica SP2 or a Zeiss LSM 710 confocal laser scanning microscope equipped with a 63x water-corrected objective was used for detection of Alexa488 (ex 488 nm/em 500-545 nm), Alexa555 (ex 561 nm/em 555-610 nm).

Microscopy and image analysis

Yellow Cameleon-based experiments were performed and analyzed as described (Behera et al., 2018). Confocal microscopy calcium imaging analyses of Arabidopsis seedlings expressing NES-YC3.60 were performed using a Nikon Eclipse Ti2 inverted microscope, equipped with a Nikon A1R+ laser scanning device (Nikon). Images were acquired by a CFI Plan Apo Lambda 20X (N.A. 0.75). Cameleon was excited with the 445 nm laser and the emissions of CFP and cpVenus were collected at 482/35 nm and 540/30 nm respectively. Pinhole was set to 35.76 μm and the images were acquired at 1024 x 1024 pixels resolution with scan speed of 0.5 frame per seconds. NIS-Elements (Nikon; http://www.nis-elements.com/) was used as a platform to control the microscope. Images were analysed using NIS-Elements to generate the ratio images and FIJI (https://imagej.net/Fiji) to quantify the fluorescence. Kymographs were generated on denoised imaged obtained with the NIS-Element Denoise.ai plugin (https://www.microscope.healthcare.nikon.com/en_EU/products/confocal-microscopes/a1hd25-a1rhd25/nis-elements-ai). By using the Fiji “Dynamic Reslice” function we created a new image, which depicts the temporal evolution of the Ca2+ signal through the spatial path previously defined.

Fluorescence emission of Alexa488 (ex 488 nm/em 500-545nm), Alexa555 (ex 561nm/em 555-610nm), YFP (ex 514nm/em 520-565nm), was detected using a 63x water objective. Images were analyzed using Fiji (Schindelin et al., 2015). Fiji was used to rotate and crop images and label the region of interest of all the roots for quantification. The proportion of cells with BFA bodies was scored manually and calculated by using Excel. BoxPlotR was used to generate the box plots (Spitzer et al., 2014).

Statistical analysis

For statistical analysis of the Ca2+ imaging data, unpaired two-tailed t-tests with Welch correction for unequal standard deviations between populations were performed using GraphPad Prism (GraphPad Prism 8 for Windows 64-bit, version 8.4.1).

For statistical analysis of the immunolocalization experiments, a logistic regression was performed to compare the presence of BFA bodies in root cells of treated versus untreated roots or wild type versus mutant. A random effect was added to the model for the experiments with multiple repeats to take into account the correlation between measurements done at the same time. The analysis was performed with the glimmix procedure from SAS (Version 9.4 of the SAS System for windows 7 64bit. Copyright 2002-2012 SAS Institute Inc. Cary, NC, USA (www.sas.com)). Maximum likelihood estimation was done with the default estimation method. A Wald-type test was performed to estimate the treatment/genotype effect on the presence of BFA bodies in the root cells.

Results

Characteristics of 1-NAA-induced Ca2+ signaling

The synthetic auxin 1-NAA interferes via a non-transcriptional pathway with the accumulation of plasma membrane proteins in Brefeldin A (BFA)-induced intracellular endosomal aggregates (hereafter referred to as BFA bodies) (Paciorek et al., 2005; Robert et al., 2010), and is most prominent at high 1-NAA concentrations (Fig. 1A,B). This effect was proposed to reflect a combination of specific and non-specific effects of 1-NAA on protein biosynthesis and endosomal aggregation (Jasik and Schmelzer, 2014; Narasimhan et al., 2021; Paponov et al., 2019). The molecular mechanisms of these effects remain unknown, but are proposed to involve non-transcriptional responses (Robert et al., 2010). Therefore, we set out to determine the involvement of auxin-induced Ca2+ signaling, a poorly understood, non-transcriptional auxin response (Dindas et al., 2018; Shih et al., 2015; Vanneste and Friml, 2013).

Fig. 1. The dose-response of BFA body formation and Ca2+ response.

Fig. 1

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(A) Immunolocalization of PIN1 in 3 day-old roots in 1/2MS (co)treated for 1h with 25μM BFA, 25μM BFA and 0.1μM 1-NAA, 25μM BFA and 1μM 1-NAA, and 25μM BFA and 10μM 1-NAA. White arrows indicate PIN1-accumulating.

(B) Quantification of the proportion of cell accumulating PIN1 in BFA bodies in (A) (For the respective treatments n=18; 22; 21; 17 roots). n is the bulked number of quantified roots from 3 independent experiments. For the box plots, significant differences (P ≤ 0.05, Wald-type test) are indicated by different lowercase letters. The central line indicates the median, the bottom and top edges of the box the interquartile range, and the box plot whiskers are plotted down to the minimum and up to the maximum value.

(C-D). Averaged and normalized NES-YC3.60 fluorescence ratios over time using wide-field illumination upon treatment with 0.1, 1 or 10μM 1-NAA. (n ≥5) respectively, means + S.D.). 1-NAA treatments were applied as indicated by the black line. The schematic root indicates the region of interest (ROI) used for measurements (C). The maximal NES-YC3.60 responses were quantified and plotted as box-and-whisker plots (D). Experimental points are plotted and their distribution represented as a box that extends from the 25th to 75th percentiles. The central line in the boxes are at the respective medians. (Unpaired Student’s t-test; * P<0.05, **P<0.01, *** P<0.001).

Using the ratiometric NES-YC3.60 (Krebs et al., 2012), we observed an instant cytosolic Ca2+ elevation in response to 10 μM 1-NAA. The onset of the Ca2+ increase started within seconds after 1-NAA application, and rapidly reached a maximum, followed by a gradual response attenuation in the presence of 1-NAA, and a rapid return to baseline levels upon 1-NAA washout (Fig. 1C). Subcellular targeting of ratiometric Ca2+ sensors revealed a rise in Ca2+ concentrations in response to 1-NAA, besides the cytosol near the plasma membrane (visualized with PM-YC3.60-Lti6b (Krebs et al., 2012)), in the endoplasmic reticulum lumen (visualized with CRT-D4ER (Bonza et al., 2013)) and in mitochondria (visualized with 4mt-YC3.60 (Loro et al., 2012)) (Supplementary Fig. S1A-D). The signals in the ER and mitochondria were slightly delayed compared to the other reporters, suggesting that these organelles may act as Ca2+ sinks for attenuation of the cytoplasmic Ca2+ signal (Resentini et al., 2021b), and is consistent with the proposal that the periplasm is the main source of auxin-induced Ca2+ (Lamport et al., 2021).

Also at lower concentrations (1 μM and 0.1 μM), 1-NAA triggered rapid Ca2+ signaling, albeit with smaller amplitudes (Fig. 1C,D), illustrating a dose-dependence of the maximal response. These analyses show that the potent inhibition of PIN1 BFA body formation by 1-NAA coincides with intense Ca2+ signaling.

The ability of auxin analogues to modulate PIN BFA body formation does not correlate with Ca2+ signaling

Different auxin analogues and anti-auxins have been shown to inhibit PIN accumulation in BFA bodies at different concentrations (Paciorek et al., 2005; Paponov et al., 2019; Robert et al., 2010; Simon et al., 2013). Therefore, we asked if this effect on trafficking would correlate with Ca2+ responses elicited by such molecules. We collected representatives of the most important classes of auxin analogues: 1-NAA, Quinclorac, 2,4-dichlorophenoxy acetic acid (2,4-D), Dicamba and Picloram, the anti-auxins α-(phenyl ethyl-2-one)-indole-3-acetic acid (PEO-IAA), Auxinole, the natural auxin Phenylacetic Acid (PAA), the presumed inactive 1-NAA analog, 2-NAA, and the non-auxinic, weak acid control Benzoic Acid (BA) (Supplementary Fig. S2). Each of these molecules was tested at 10μM to allow for comparison of the effects.

Consistently with previous reports (Paponov et al., 2019; Robert et al., 2010), 1-NAA, 2-NAA, and both anti-auxins strongly inhibited PIN1 accumulation in BFA bodies (Fig. 2A,B). Of all other tested molecules, only 2,4-D caused a significant reduction in the proportion of cells with PIN1 in BFA bodies (Fig. 2A,B). It is known that 2,4-D exerts stronger trafficking effects at higher concentrations (Simon et al., 2013). Similar effects of these chemicals were observed for PIN2 accumulation in BFA bodies (Supplementary Fig. S3A,B).

Fig. 2. Distinct effects of different auxin analogs on PIN1 BFA body formation.

Fig. 2

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(A) Analysis of PIN1 internalization through whole-mount immunolocalization in 3 day-old Col-0 seedling root meristems. Seedlings were treated with 25μM BFA and 10μM of 1-NAA, 2-NAA, Quinclorac, 2,4-D, Phenylacetic acid, Benzoic acid, Dicamba, Picloram, PEO-IAA, Auxinole. Scale bar=10 μm.

(B) Quantification of the proportion of cell accumulating PIN1 in BFA bodies for different co-treatments described in (A) (n=18; 18; 16; 13; 17; 14; 14; 11; 14; 16; 17 roots). n is the bulked number of quantified roots from 3 independent experiments. For the box plots, significant differences (P ≤ 0.05, Wald-type test) are indicated by different lowercase letters. The central line indicates the median, the bottom and top edges of the box the interquartile range, and the box plot whiskers are plotted down to the minimum and up to the maximum value.

Next, we analyzed the effects of pulse of the respective molecules on the Ca2+ response. Surprisingly, the “inactive” 1-NAA analog, 2-NAA triggered a Ca2+ response that was a bit slower and had a lower amplitude than 1-NAA (Fig. 3A,B), indicating 2-NAA is not completely inactive (Fig. 3A,B). In contrast to 1-NAA, the acute Ca2+ response to 2,4-D did not return to basal levels after wash-out (Fig. 3D). This could be a consequence of 2,4-D being a poor substrate for efflux machinery and being trapped inside the cell after uptake (Delbarre et al., 1996). In contrast, neither quinclorac, PAA, BA, dicamba, picloram, PEO-IAA nor auxinole activated an acute Ca2+ response (Fig. 3C, E-H). The slow linear induction of Ca2+ signalling during PAA treatment is somewhat surprising, as this natural auxin analog is also promiscuously sensed via TIR1/AFBs (Lee et al., 2014; Sugawara et al., 2015).

Fig. 3. Distinct Ca2+ dynamics in response to different auxin analogs.

Fig. 3

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(A-J) Averaged and normalized NES-YC3.60 fluorescence ratios over time using wide-field illumination upon treatment with 10μM 1-NAA (A), 2-NAA (B), Quinclorac (C), 2,4-D (D), Phenylacetic acid (E), Benzoic acid (F), Dicamba (G), Picloram (H), PEO-IAA (I) and Auxinole (J). Dashed lines indicate the interval of pulse treatment. (K) Quantification of the maximal NES-YC3.60 responses plotted as box-and-whisker plots (n=5 for each treatment described in (A-J), means + S.D. (STATISTICS)

Together with the BFA body formation data, these data suggests that the ability of auxin analogues to inhibit PIN BFA body formation roughly correlates with their ability to induce a Ca2+ response. However, this correlation does not hold true for the anti-auxins, as none of them induced an acute Ca2+ response (Fig. 3I-K).

1-NAA-induced Ca2+ signaling depends on TIR1/AFB-CNGC14

While we observed a correlation between the ability to inhibit PIN BFA body formation and Ca2+ signaling for the tested auxin analogues (Fig. 2B, 3K), this correlation does not hold true for the endogenous auxin IAA: IAA triggers rapid Ca2+ signals (Dindas et al., 2018), but does not strongly interfere with BFA body formation (Narasimhan et al., 2021; Paponov et al., 2019). This could signify that 1-NAA employs a Ca2+ signaling mechanism, that is independent of IAA-induced Ca2+ signaling, to inhibit BFA body formation.

Therefore, we tested the involvement of IAA-induced Ca2+ signaling components in 1-NAA-induced Ca2+ signaling(Dindas et al., 2018; Li et al., 2021; Shih et al., 2015). We transformed the ratiometric NES-YC3.60 construct into tir1afb2afb3 and cngc14-1 mutants. Transformants with expression levels similar to our WT control were used to monitor NAA-induced Ca2+ signals (Fig. 4A). The 1-NAA-induced Ca2+ response was strongly reduced in the tir1afb2,3 triple mutants and was completely absent from cngc14-1 (Fig. 4B,C). This corroborates that 1-NAA activates Ca2+ signaling via a TIR1/AFB-CNGC14-dependent mechanism, and is consistent with the measured reductions of extracellular Ca2+ fluxes in these mutants (Dindas et al., 2018).

Transcriptional reporters and single cell expression analyses indicate that CNCG14 expression is largely restricted to the root epidermis (Fig. 4D)(Brost et al., 2019; Wendrich et al., 2020). Therefore, we explored the contributions of the different root tissues to 1-NAA-induced Ca2+ signaling by also analyzing the NES-YC3.60 signal via confocal scanning microscopy (Fig. 4E), instead of wide-field illumination (Fig. 4B). To visualualize the response dynamics in all tissues we defined a single line across the root segment of interest and plotted the NES-YC3.60 ratios over time to generate a kymograph (Fig. 4F). In contrast to the epidermis-specific expression of CNGC14, the kymograph revealed a near simultaneous 1-NAA-induced Ca2+ response in all root tissues (Fig. 4F). What is more, the 1-NAA-induced Ca2+ response was CNGC14-dependent in all tissues (Fig. 4F). This indicates that CNGC14 indirectly controls auxin-induced Ca2+ responses in all tissues of the root meristem, possibly involving complex signal propagation and amplification mechanisms based on plasmodesmata and electrical signaling (Fichman et al., 2021; Nguyen et al., 2018; Resentini et al., 2021a). Together, these data demonstrate that 1-NAA-induced Ca2+ signaling depends on TIR1/AFB and CNGC14 activities.

Therefore, we could now address the requirement for 1-NAA-induced Ca2+ signalling in the inhibition of PIN1 BFA body formation. We tested a series of single, double and triple tir1/afb mutants, as well as three cngc14 alleles and double and triple mutant combination with mutants in related CNGCs with demonstrated functional redundancies (CNGC6, and CNGC9) (Brost et al., 2019). All mutants formed BFA bodies (Fig. 5A,C; Supplementary Fig. S4A-C), that could be inhibited by 1-NAA (Fig. 5B,C; Supplementary Fig. S4A-C), suggesting that the TIR1/AFB-CNGC14 module is not required for this effect of 1-NAA. We also tested the sensitivity of the cngc2-3 mutant, which was recently found to be defective in IAA-induced Ca2+ signaling (Chakraborty et al., 2021). Similarly to tir1/afb and cngc14 mutants, the 1-NAA sensitivity of BFA body formation was not changed in cngc2-3 compared to WT (Supplementary Fig. 5).

Fig. 5. Analysis of 1-NAA sensitive BFA body formation in tir1/afb and cngc14 mutants.

Fig. 5

Open in a new tab

(A-B) Immunolocalization of PIN1 in 3 day-old roots treated with BFA (A) or co-treated for BFA and 1-NAA

(B) in wild type, tir1afb1afb3, tir1afb2afb3, cngc14-1 and cngc14-2. White arrows indicate PIN1-accumulating BFA bodies. Scale bar is 20 μm.

(C) Quantification of the proportion of cells that accumulate PIN1 in BFA bodies visualized by immunolocalization as illustrated in (A) and (B) for wild type (nA =11; nB =11), tir1afb1afb3 (nA =10; nB =12), tir1afb2afb3 (nA =11; nB =11), cngc14-1 (nA =11; nB =12) and cngc14-2 (nA =11; nB =12). n is the pooled number of roots derived from three independent replicates.

Jointly, these data demonstrate that 1-NAA inhibits BFA body formation independently of TIR1/AFB-CNGC14-mediated Ca2+ signaling.

For the box plots, significant differences (P ≤ 0.05, Wald-type test) are indicated by different lowercase letters. The central line indicates the median, the bottom and top edges of the box the interquartile range, and the box plot whiskers are plotted down to the minimum and up to the maximum value.

Discussion

CNGC14 is required for 1-NAA-induced Ca2+ signaling in all root tissues

In root hairs, CNGC14 acts partially redundant with other CNGCs to generate and maintain tip-focused Ca2+ oscillations (Brost et al., 2019; Zhang et al., 2017). In the root meristem, however, a single mutation in CNGC14 is sufficient to abrogate IAA-induced Ca2+ signaling (Dindas et al., 2018; Shih et al., 2015). Similarly, we found that cngc14 mutants are defective in 1-NAA-induced Ca2+ signaling, suggesting that 1-NAA and IAA activate similar Ca2+ signaling mechanisms.

In performing our analyses, we found that the 1-NAA-induced Ca2+ response is near instantaneous in all tissues of the root. Notably, single cell expression datasets showed that CNGC14 is predominantly expressed in the epidermis, and is possibly absent in the inner root tissues. Yet, the entire 1-NAA-induced Ca2+ response was defective in the cgnc14 mutant. This discrepancy between the expression domain and the observed effects suggests non-cell autonomous coordination of auxin-induced Ca2+ signaling in the root meristem. Such systemic Ca2+ responses are emerging as a prevalent theme in plant Ca2+ signaling, in which different Ca2+ channel types, in conjunction with electrical signals, reactive oxygen species and plasmodesmata, are involved in initiation and propagation of intercellular Ca2+ signaling (Choi et al., 2014; Fichman and Mittler, 2020; Fichman et al., 2021; Resentini et al., 2021a; Toyota et al., 2018). Intercellular coordination of auxin-induced Ca2+ via similar mechanism can be expected, given the requirement of coordination of growth of different tissues during gravitropic bending. Moreover, ROS production in response to a mechanical stimulus has been connected to Ca2+ signaling in the root (Monshausen et al., 2009). Plasmodesmata are also emerging as regulators of auxin distribution and activity (Gao et al., 2020; Mellor et al., 2020; Sager et al., 2020), indicating that these auxin-regulated plasmodesmata could modulate intracellular ROS movement for amplification of auxin-regulated Ca2+ signaling.

Does AFB1 contribute to auxin-induced Ca2+ signaling?

Recent efforts identified TIR1/AFBs as auxin sensors for auxin-induced Ca2+ signaling. This was demonstrated by the induction of rapid Ca2+ signaling by the orthogonal activation of an engineered TIR1 variant (ccvTIR1) (Li et al., 2021). In contrast to tir1 single mutants, auxin-induced Ca2+ signaling is strongly reduced tir1afb2,3 triple mutants (Dindas et al., 2018; Monshausen et al., 2011), demonstrating functional redundancy among TIR1/AFBs. The fact that we could still detect an important auxin-induced Ca2+ signaling in tir1afb2,3 triple mutants, but not in cngc14 (Fig. 4B,C), indicates that additional TIR1/AFBs are involved in auxin-induced Ca2+ signaling.

Two of the used auxin analogues, picloram and quinclorac, have a strong biochemical preferences for AFB4/AFB5 over TIR1-type AFBs (Calderon Villalobos et al., 2012; Lee et al., 2014; Prigge et al., 2016), suggesting they selectively activate AFB4 and AFB5. As neither of these compounds activated a detectable Ca2+ response (Fig. 3), we propose that these AFBs have minor, if any, roles in auxin-induced Ca2+ signaling. By excluding AFB4 and AFB5, we suggest that AFB1 auxin sensing activities could explain the majority of residual auxin-induced Ca2+ signaling in tir1afb2,3 triple mutants. Consistently with a role in membrane associated auxin signaling, AFB1 is most closely related to TIR1, has a non-nuclear localization (Prigge et al., 2020), and was recently highlighted as an auxin sensor for non-transcriptional membrane depolarization and initiation of root growth inhibition (Serre et al., 2021).

1-NAA inhibits BFA body formation independently of TIR1/AFB-CNGC14-dependent Ca2+ signaling

Auxin-mediated regulation of trafficking is considered as an important aspect of auxin’s self-regulating properties in plant growth and development. However, the underlying molecular mechanisms remain largely unclear. Synthetic auxins, such as 1-NAA, display non-specific effects on the endomembrane system, such as a strong interference with aggregation of endosomes in BFA bodies (Narasimhan et al., 2021; Paponov et al., 2019). This effect was reported to be very fast, not requiring transcriptional changes and independent of canonical auxin signaling (Robert et al., 2010). Given the immediacy of the response, we explored if Ca2+ signaling could be involved. We observed some correlation between the ability to inhibit PIN1 accumulation in BFA bodies and to stimulate Ca2+ signaling at high 1-NAA concentrations (Fig. 1) and for different auxin analogs (Fig. 2, 3). Interestingly, the presumed “inactive” 2-NAA (Paponov et al., 2019), also triggered a pronounced Ca2+ response, possibly via its weak affinity for TIR1 (Lee et al., 2014), indicating 2-NAA is not an ideal negative control for fast auxinic responses. However, this correlation did not hold true for anti-auxins (Fig. 2,3), and IAA (Dindas et al., 2018; Paponov et al., 2019), indicating that different auxin signaling pathways could be involved.

We established that 1-NAA activates Ca2+ responses at the plasma membrane via a TIR1/AFB-CNGC14-dependent mechanism, similarly as was previously described for the endogenous auxin IAA (Dindas et al., 2018; Shih et al., 2015). This shows that 1-NAA-induced Ca2+ signaling is a specific auxinic response. However, this genetic disruption of 1-NAA-induced Ca2+ signaling in tir1afb2,3 and cngc14 mutants did not affect 1-NAA’s ability to inhibit PIN accumulation in BFA bodies. Moreover, the cncg2 mutant that is defective in auxin signaling and homeostasis {Chakraborty, 2021 #929}, also did not alter the 1-NAA sensitivity of BFA body formation. These data unequivocally show that BFA body formation is not inhibited by auxin-induced Ca2+ signaling, and further support the notion that the synthetic auxin 1-NAA differentially affects the endomembrane system compared to IAA. It does, however, not exclude roles for TIR1/AFB-CNGC14-mediated Ca2+ signaling in the auxin-regulated vacuolar trafficking of PIN proteins, that depends on TIR1/AFB function (Baster et al., 2013), or PIN polarization (Mazur et al., 2020a; Mazur et al., 2020b; Prat et al., 2018; Sauer et al., 2006). The use of genetic tools will be imperative to resolve which parts of the responses to auxin analogs can be attributed to a particular auxin signaling mechanism.

Despite its limitations, 1-NAA’s antagonism towards endomembrane aggregation and BFA body formation has been a convenient pharmacological tool to identify and characterize new regulators of endomembrane trafficking (Feraru et al., 2012; Tanaka et al., 2009; Zhang et al., 2020).

Supplementary Material

Suppl Figures

Highlight.

The effect of synthetic auxin analogs on non-transcriptional responses is poorly understood. Here, we characterize auxin analog-induced Ca2+ signaling, and show that this is independent of inhibition of endosomal aggregation.

Acknowledgements

We thank Joerg Kudla (WWU Munster, Germany), Petra Dietrich (F.A. University of Erlangen-Nurnberg, Germany) for sharing published materials, and NASC for providing seeds. We thank Veronique Storme for help with the statistical analyses. This work was supported by grants of the China Scholarschip Council (CSC) (to R.W. and J.C.), Fonds Wetenschappelijk Onderzoek (FWO) (to T.B. and S.V.), the special research fund Ghent University (to E.H.), the European Research Council (ERC) (T-Rex project number 682436 to D.V.D and J.F.), the Deutsche Forschungsgemeinschaft (DFG) through Grants within FOR964 (M.K. and K.S.), Piano di Sviluppo di Ateneo 2019 (University of Milan) (to A.C.), and by a PhD fellowship from the University of Milan (to M.G.). Part of the imaging analyses were carried out at NOLIMITS, an advanced imaging facility established by the University of Milan.

Footnotes

Author Contributions

R.W and E.H. performed immunolocalizations and subsequent data analyses according to J.F. methodology and resources. E.H. and M.G. performed the Ca2+ reporters experiments according to M.K., K.S. and A.C. methodology. S.V. and A.C. conceived the original screening and research plans and wrote the manuscript with contributions of all the authors. All authors were involved in discussing and analyzing the data. S.V. and A.C agree to serve as the authors responsible for contact and ensure communication.

Conflict of interest

The authors declare no conflict of interest

Data Availability

All data generated or using in the submitted article are available from the corresponding authors upon request.

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