https://orcid.org/0000-0001-5449-6492
Abstract
The phytohormone auxin governs various developmental processes in plants including vascular formation. Auxin transport and biosynthesis are important factors in determining auxin distribution in tissues. Although the role of auxin transport in vein pattern formation is widely recognized, that of auxin biosynthesis in vascular development is poorly understood. Heterodimer complexes comprising two basic helix–loop–helix protein families, LONESOME HIGHWAY (LHW) and TARGET OF MONOPTEROS5 (TMO5)/TMO5-LIKE1 (T5L1), are master transcriptional regulators of the initial process of vascular development. The LHW–TMO5/T5L1 dimers regulate vascular initial cell production, vascular cell proliferation and xylem fate determination in the embryo and root apical meristem (RAM). In this study, we investigated the function of local auxin biosynthesis in initial vascular development in RAM. Results showed that LHW–T5L1 upregulated the expression of YUCCA4 (YUC4), a key auxin biosynthesis gene. The expression of YUC4 was essential for promoting xylem differentiation and vascular cell proliferation in RAM. Conversely, auxin biosynthesis was required for maintaining the expression levels of LHW, TMO5/T5L1 and their targets. Our results suggest that local auxin biosynthesis forms a positive feedback loop for fine-tuning the level of LHW–TMO5/T5L1, which is necessary for initiating vascular development.
Introduction
The formation of vascular initial cells first occurs in the lower middle part of the embryo at the globular stage. Recent studies demonstrate that heterodimer complexes comprising LONESOME HIGHWAY (LHW) and TARGET OF MONOPTEROS5 (TMO5)/TMO5-LIKE1 (T5L1) regulate cell divisions for the generation of vascular initial cells in an embryo, and upregulate cytokinin biosynthesis, which contributes to the proliferation of vascular stem cells in embryos and root apical meristem (RAM) (Ohashi-Ito and Bergmann 2007, De Rybel et al. 2013, Ohashi-Ito et al. 2013, De Rybel et al. 2014, Ohashi-Ito et al. 2014). The LHW–TMO5/T5L1 dimers also establish xylem cell identity (Katayama et al. 2015). Thus, LHW–TMO5/T5L1 are considered as key complexes regulating the initial process of vascular development in an embryo and RAM. In addition to LHW–TMO5/T5L1, auxin is another key regulator of initial vascular development (Cano-Delgado et al. 2010). Therefore, it is reasonable to speculate that LHW–TMO5/T5L1 and auxin signaling pathways are associated with each other in the initial process of vascular development. Indeed, TMO5 is a target of MONOPTEROS/AUXIN RESPONSE FACTOR5 (MP/ARF5; Schlereth et al. 2010) and is auxin-inducible. However, the functional relationship between LHW–TMO5/T5L1 and auxin and the role of transcription factor-dependent auxin biosynthesis in the initiation of vascular development are poorly understood. We found that YUCCA4 (YUC4) was listed as a candidate downstream gene of LHW–T5L1 in a transcriptome analysis of LHW–T5L1 overexpressing cells (Ohashi-Ito et al. 2014). Because YUC4 is well known as a key auxin biosynthetic enzyme (Cheng et al. 2006, Cheng et al. 2007), we here investigated the functional relationship between LHW–TMO5/T5L1 and auxin in initial vascular development, placing YUC4 at the heart of the analysis.
Results
To elucidate the role of YUC4 in vascular development, we first determined whether the expression of YUC4 was regulated by LHW and T5L1 in Arabidopsis culture cells simultaneously overexpressing LHW and T5L1-GFP under the control of an estrogen-inducible promoter. The expression of YUC4 was conspicuously upregulated >70-fold at 24 h after the addition of estrogen, with high expression levels at 48 and 72 h (Fig. 1A). Next, we performed chromatin immunoprecipitation (ChIP)-PCR assay using culture cells, which harboring the estrogen-inducible LHW/T5L1-GFP or the estrogen-inducible YFP. The estrogen-inducible YFP line was used as a control, and immunoprecipitates of this sample were the background level. Results of ChIP-PCR analysis showed that YUC4 promoter fragments, but not coding sequence fragments, were enriched in the immunoprecipitates of LHW and T5L1-GFP overexpressing cells (Fig. 1B, C), suggesting that T5L1 directly binds to the YUC4 promoter in the presence of LHW. Next, we generated plants harboring both pYUC4::GUS and estrogen-inducible LHW and T5L1-GFP constructs. Although GUS activity driven by the YUC4 promoter was restricted to the vascular region of the root in the absence of estrogen, ectopic GUS activity was observed in various root cells when the expression of LHW and T5L1 was induced after the addition of estrogen (Fig. 1D, E), indicating that the combination of LHW and T5L1 can induce YUC4 expression. Next, we examined the expression patterns of pYUC4::nuclear-localized YFP (nYFP) in wild-type and lhw roots. The nYFP signal driven by the YUC4 promoter was observed in metaxylem precursor cells in wild-type RAM (Fig. 1F, H). Because LHW is broadly expressed in RAM and TMO5 and T5L1 are expressed in xylem precursor cells, the LHW–T5L1 heterodimer is considered to function in xylem precursor cells in RAM (Ohashi-Ito and Bergmann 2007, De Rybel et al. 2013). In RAM, therefore, the YUC4 expressing cells overlapped with LHW–T5L1 functioning cells. In lhw RAM, the number of YUC4 expressing cells was dramatically decreased (Fig. 1G, I). Together, these results imply that LHW–T5L1 directly regulates YUC4 expression in the RAM.
LHW–T5L1 regulates YUC4 expression. (A) Quantitative reverse transcription-PCR (RT-PCR) analysis of YUC4 expression levels in cultured cells harboring estrogen-inducible LHW and T5L1-GFP with (black) or without (light blue) estrogen. Data represent the mean ± standard deviation (SD). The mean is the average of three independent biological replicates. Significant differences were determined using Student’s t-test and are indicated with asterisks (***P < 0.001). (B) Schematic representation of the YUC4 gene. Black boxes indicate exons, and black bars indicate positions of primers used in the ChIP-PCR experiment. (C) Quantitative ChIP-PCR analysis using cells including estrogen-inducible LHW and T5L1-GFP (light blue, black), and cells including estrogen-inducible YFP (yellow, gray) with estrogen (black, gray) and without estrogen (light blue, yellow). Data represent mean ± SD. Significant differences between LHW–T5L1 Est− vs. YFP Est− and LHW–T5L1 Est+ vs. YFP Est+ were determined using Student’s t-test and are indicated with asterisks (**P < 0.01). The mean is the average of three independent biological replicates. (D, E) Expression patterns of pYUC4::GUS in the differentiation zone of roots of 8-day-old plants harboring estrogen-inducible LHW–T5L1 treated with (E) or without (D) estrogen for 24 h. Scale bar = 100 µm. (F–I) Expression patterns of pYUC4::nYFP in 5-day-old wild-type (F) and lhw (G) root tips. Cross-sections of (F) and (G) are shown in (H) and (I), respectively. Asterisks indicate protoxylem cells. Scale bar = 100 µm.
Next, we characterized the phenotype of a loss-of-function yuc4 mutant, which had a T-DNA insertion in the 5′-UTR region of YUC4 and the transcript level of YUC4 in yuc4 was greatly reduced compared with that of the wild type (Supplementary Fig. S1). In yuc4 plants, the root vasculature showed a diarch pattern, which consists of two protoxylem and protophloem poles, with normally differentiated protoxylem and metaxylem vessels (Fig. 2A, B, F, G), similar to that observed in wild-type Arabidopsis roots. However, yuc4 roots contained fewer vascular stem cells (procambial cells) than wild-type roots, although the number of xylem and phloem cells was similar to that in wild-type roots (Fig. 2J, K). To further examine the role of YUC4 in vascular development, we generated lhw yuc4 double mutant plants, because the lhw mutant exhibited major defects in vascular development and thus it was suitable for the analysis of vascular development as a sensitive background. The phenotype of about one-third of the lhw yuc4 roots was similar to that of lhw roots, such as a monarch pattern vasculature; however, in the remaining two-thirds of the lhw yuc4 roots, xylem vessel differentiation was partially arrested because of the delay of differentiation, which was a novel phenotype (Fig. 2C-E, H, I). This result implies that LHW is not the only regulator of YUC4 and other factors including LHW-family proteins may also regulate YUC4 expression. The arrest of xylem vessel differentiation in lhw yuc4 mutant roots was complemented by the expression of pYUC4::YUC4 (Fig. 2L). To determine whether YUC4 expression in RAM is required for proper vessel differentiation, we overexpressed YUC4 under the control of a RAM-specific promoter RGFR1 (Shinohara et al. 2016) in the lhw yuc4 double mutant background. The expression of pRGFR1::YUC4 complemented the arrested differentiation phenotype of xylem vessels (Fig. 2M). To further examine the importance of auxin biosynthesis in vascular development, we observed plants that are treated with kynurenine, an inhibitor of auxin synthesis. The width of a stele in RAM was significantly decreased in kynurenine-treated plants (Supplementary Fig. S2A–C). In addition, delay and partial arrest of xylem differentiation frequently occurred in kynurenine-treated lhw roots (Supplementary Fig. S2D, E). These results suggest that auxin synthesis is required for promoting vascular cell proliferation and proper xylem differentiation.
Phenotypes observed in the vascular region of yuc4 roots. (A–E) Images of vascular tissue in 7-day-old wild-type (A), yuc4 (B), lhw (C) and lhw yuc4 (D, E) differentiation zone of roots. Scale bar = 50 µm. The fraction of samples showing similar patterns is presented. (F–I) Cross-sections of the vascular region in 7-day-old wild-type (F), yuc4 (G), lhw (H) and lhw yuc4 (I) differentiation zone of roots. Procambial cells, xylem precursor cells and phloem cells are indicated in yellow, green and blue, respectively. Cell types were judged by their morphological features. Scale bar = 20 µm. (J) Number of vascular cells in wild-type, yuc4, lhw and lhw yuc4 roots. Data represent mean ± SD; n = 14 (wild-type), 13 (yuc4), 12 (lhw) and 8 (lhw yuc4). The means, indicated by different letter superscripts in each column, were significantly different according to one-way ANOVA with the Tukey–Kramer post hoc test (P < 0.01). (K) Number of xylem, phloem and vascular stem cells in wild-type (black) and yuc4 (gray) roots. Data represent mean ± SD; n = 13. **P < 0.01 (Student’s t-test). (L, M) Images of vascular tissue in 7-day-old lhw yuc4 differentiation zone of roots harboring pYUC4::YUC4 (L) or pRGFR1::YUC4 (M). Scale bar = 50 µm. The fraction of samples showing similar patterns is presented.
Indole-3-acetic acid (IAA), a major auxin, is synthesized from tryptophan in two steps, and YUC4 is one of the enzymes, which catalyze the second step in this pathway (Mashiguchi et al. 2011). To determine whether the LHW–T5L1 dimer promotes auxin biosynthesis, we monitored changes in IAA concentration after the induction of LHW and T5L1-GFP. The amount of IAA started to increase at 72 h after the induction of LHW–T5L1, and was accumulated to high levels at 96 h, whereas the amount of IAA did not change when LHW–T5L1 overexpression was not induced (Fig. 3A). This result confirms that LHW–T5L1 can activate auxin biosynthesis.
Auxin biosynthesis maintains the expression level of LHW–TMO5-related genes. (A) Concentration of IAA in cells harboring estrogen-inducible LHW and T5L1-GFP with estrogen (black) and without estrogen (gray). Data represent mean ± SD (n = 5). (B, C) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (B), and LOG3, LOG4 and ACL5 (C), in 7-day-old root tips treated with 100 µM kynurenine (Kyn) or water (control) for 0, 24 and 48 h. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates. (D, E) Quantitative RT-PCR analysis of expression levels of LHW, TMO5 and T5L1 (D) and LOG3, LOG4 and ACL5 (E) in 7-day-old wild-type and yuc4 root tips. Data represent mean ± SD (*P < 0.05, **P < 0.01; Student’s t-test). The mean is the average of three independent biological replicates.
The expression of TMO5 and T5L1 is under the regulation of auxin because these genes are targets of MP/ARF5 (Schlereth et al. 2010). We hypothesized that auxin biosynthesis promoted by LHW–T5L1 is needed for maintaining the expression level of LHW and T5L1. To test this hypothesis, we first examined the expression level of LHW, TMO5 and T5L1 and their target genes such as LOG3, LOG4 and ACL5 in plants treated with kynurenine for 48 h. After starting the kynurenine treatment, the fluorescence of DR5, an auxin response marker (Heisler et al. 2005), was decreased at 3 h, very weak at 6 h and almost undetectable at 24 h (Fig. 4G–L). The kynurenine treatment caused significant reduction in the expression level of LHW, TMO5 and T5L1 and their target genes, LOG3, LOG4 and ACL5 (Fig. 3B, C). Especially, the T5L1 expression level was greatly reduced. We also monitored the temporal expression pattern of pT5L1-nYFP after the inhibition of auxin biosynthesis with kynurenine. The nYFP signal greatly decreased at 3 h after kynurenine treatment and was completely abolished by 24 h (Fig. 4A–F). These results strongly support our hypothesis of a positive feedback loop in which auxin biosynthesis promotes the expression of LHW, TMO5 and T5L1, which in turn promotes the expression of their target genes. Gene expression analysis indicated that LOG4 and ACL5 were significantly downregulated in yuc4 root tips compared with wild-type root tips (Fig. 3E), although no significant difference was detected in the expression levels of LHW, TMO5 and T5L1 between wild-type and yuc4 root tips (Fig. 3D). Although these data support our hypothesis at least partially, they also indicate that YUC4 may not be the only player of auxin biosynthesis in LHW–TMO5/T5L1-dependent vascular events. Previous studies showed abnormal vein patterns in leaves of quadruple mutants of yuc genes, including yuc4 (Cheng et al. 2006, Cheng et al. 2007), suggesting that other YUC genes are also involved in initial vascular development.
The inhibition of auxin biosynthesis changes gene expression patterns in vascular cells. (A–R) Expression patterns of T5L1 (A–F), DR5 (G–L) and TCSn (M–R) in the RAM of 5-day-old plants treated with or without 100 µM kynurenine (Kyn) for 3, 6 and 24 h. Roots were stained with propidium iodide (purple). Scale bar = 100 µm.
In both auxin-related experiments, auxin reduction decreased LOG gene expression. Therefore, we examined the expression of a cytokinin response marker, TCSn (Zurcher et al. 2013), as an output of LHW–T5L1 activity. The TCSn signal decreased gradually from 6 to 24 h after the start of the kynurenine treatment, as expected (Fig. 4M–R). This result also suggests that the inhibition of auxin biosynthesis causes the decrease of the LHW–TMO5/T5L1 activity because we previously showed that the TCSn signal is absent at the vascular region in lhw and log3 log4 mutant root tips (Ohashi-Ito et al. 2014).
Discussion
In this study, we showed that LHW–T5L1, which is a key regulator of vascular development in RAM, promotes auxin biosynthesis. It is shown that auxin biosynthesis in RAM is essential for metaxylem differentiation (Ursache et al. 2014). Our results also showed that auxin biosynthesis in RAM promoted xylem differentiation. Thus, the control of xylem differentiation driven by auxin biosynthesis may occur quite early in vascular development. It is reported that LHW–TMO5/T5L1 positively regulate the biosynthesis of two phytohormones or phytohormone-like molecules, in addition to auxin. One of these molecules is cytokinin, which spreads into neighboring cells and induces the division of vascular cells, whereas the other is thermospermine, which functions in the negative feedback loop affecting LHW–TMO5/T5L1 (De Rybel et al. 2014, Ohashi-Ito et al. 2014, Katayama et al. 2015, Vera-Sirera et al. 2015). Thus, LHW–TMO5/T5L1 regulate the synthesis of at least three phytohormones or phytohormone-like molecules. Among these, auxin and thermospermine act in a cell-autonomous manner, whereas cytokinin acts in a noncell-autonomous manner. A combination of these two types of signal molecules may be important for generating a new tissue to spatiotemporally coordinate cell division and differentiation.
Our results suggest that LHW–T5L1 and auxin biosynthesis are tightly related in the initial process of vascular development, and these two factors mutually regulate each other. The LHW–T5L1 dimer upregulates YUC4 and activates auxin biosynthesis. Conversely, auxin promotes the expression of LHW, TMO5 and T5L1. Therefore, it is likely that LHW–T5L1/TMO5 and YUC4-dependent auxin biosynthesis form a positive feedback loop. Although it is well known that auxin flow is regulated through a positive feedback loop (Linh et al. 2018), the concept that auxin biosynthesis is a component of a positive feedback loop is novel in vascular development. The LHW–T5L1/TMO5 dimers also form a negative feedback loop through the upregulation of ACL5 and SACL family genes, which encode a thermospermine synthase and bHLH proteins, respectively (Katayama et al. 2015, Vera-Sirera et al. 2015). The synthesized thermospermine promotes the production of SACL proteins, which interact with LHW and block the formation of the active LHW–T5L1/TMO5 heterodimers. This negative feedback regulation suppresses the function of LHW–T5L1/TMO5 at the protein level, whereas the positive feedback regulation, comprising auxin biosynthesis, promotes the function of LHW–T5L1/TMO5 at the transcript level. Although mechanisms underlying the positive and negative feedback regulations of LHW–T5L1/TMO5 are different, both these regulations likely control the output of LHW–T5L1/TMO5 as a whole. In other words, the balance between positive and negative feedback regulations may be responsible for fine-tuning the output of LHW–T5L1/TMO5 (Supplementary Fig. S3). We note, however, that other YUC4-dependent transcription factor than LHW–TMO5/T5L1 may be also involved in the initial process of vascular development, because the number of vascular cells decreased in yuc4, despite the fact that expression levels of LHW, TMO5 and T5L1 were not significantly changed between wild type and yuc4. In this case, this unknown factor might be reduced in yuc4 mutants, resulting in a decrease in the expression level of LOG4, which promotes cell division by producing cytokinin. ATHB8 is a possible candidate of this factor because ATHB8 is auxin-inducible (Ursache et al. 2014) and regulates ACL5 (Baima et al. 2014), which expression was lower in yuc4.
Previous studies showed that local auxin biosynthesis plays crucial roles in many aspects of plant development such as the formation of floral organs, the apical–basal axis formation in embryos and the maintenance of root stem cell niche (Cheng et al. 2006, Zhao 2008, Robert et al. 2013, Brumos et al. 2018). In some of these processes, local auxin biosynthesis contributes to generating auxin maximum. Our previous study showed that lateral root primordia in lhw mutants failed to confine auxin maximum in vascular cells and the quiescent center (Ohashi-Ito et al. 2013). Therefore, LHW–TMO5/T5L1-mediated local auxin biosynthesis and positive feedback regulations may contribute to the confined auxin maximum formation during vascular development.
In summary, we propose a model in which auxin biosynthesis is activated in the initial process of vascular development, and forms a positive feedback loop to control the level of LHW–T5L1/TMO5. The level of LHW–T5L1/TMO5 is also regulated by a negative feedback loop consisting of LHW–T5L1/TMO5, ACL5 and SACLs. Because initial vascular development, including the proliferation of vascular stem cells and determination of xylem cell identity, is dominated by LHW–T5L1/TMO5, it appears that adjusting the output of LHW–T5L1/TMO5 appropriately through the double feedback regulations is critical for the coordinated development of vascular tissues.
Materials and Methods
Plant materials and growth conditions
Arabidopsis thaliana accession Columbia, yuc4 (SALK_047083C), lhw (SALK_079402), DR5 rev::3x VENUS-N7 (Heisler et al. 2005) and TCSn::GFP (Zurcher et al. 2013) were used. Seeds were sown on half-strength Murashige and Skoog (1/2 MS) agar plates containing 1% sucrose, 0.8% agar and appropriate antibiotics (MS plates), incubated at 4°C for 2–3 d, and then moved to an incubator for growth under continuous light at 22°C. For kynurenine treatment, plants were grown on MS plates for 5 d and then transferred onto MS plates including 100 µM kynurenine.
Arabidopsis culture cells harboring estrogen-inducible LHW and T5L1-GFP established in the previous study were used (Ohashi-Ito et al. 2014). Estrogen-inducible YFP line was established according to previously described methods (Ohashi-Ito et al. 2010). To induce LHW and T5L1-GFP, or YFP expression, 1 ml of MS medium containing 0.5 µM estradiol was supplemented to a 0.5 ml aliquot of 10-day-old transformant cell culture and then cultured on a rotary shaker at 124 rpm at 22°C in the dark.
DNA manipulation
The promoter sequences of YUC4 (2.2 kb), RGFR1 (2.5 kb) and T5L1 (3.0 kb) were amplified by PCR and cloned into pENTR/D-TOPO cloning vector (Thermo Fisher Scientific, Waltham, MA, USA) to generate entry vectors. The promoter sequences were integrated into pBGYN vector (Kubo et al. 2005) or pGWB434 vector, which included intron-GUS for promoter analysis, and was gift from T. Nakagawa, using LR Clonase II Enzyme Mix (Life Technologies). The coding sequence of YUC4 was amplified by PCR with YUC4cds-AscI-F and YUC4cds-AscI-R primers and digested with AscI restriction enzyme (New England Biolabs, Beverly, MA, USA). This DNA fragment was ligated with AscI-digested entry vectors to generate pYUC4::YUC4 or pRGFR1::YUC4, which were transferred into pGWB1 vector (Nakagawa et al. 2007) using LR Clonase II Enzyme Mix. Primers used in this study are listed in Supplementary Table S1.
RNA isolation and quantitative RT-PCR
Total RNA isolation and cDNA synthesis were performed according to previously described methods (Ohashi-Ito et al. 2010). Quantitative RT-PCR was performed using LightCycler480 Probes Master (Roche Diagnostics, Mannheim, Germany) with sets of Universal ProbeLibrary probes (Roche) and primers listed in Supplementary Table S1 on a LightCycler480 instrument II (Roche) according to the manufacturer’s protocol. Standard curves for each primer set were created by the second derivative maximum method using 10-fold serial dilutions of cDNA obtained from one of the samples. Expression levels of each gene calculated from the standard curves were converted to expression ratio relative to those of the UBQ10 reference gene.
Chromatin immunoprecipitation-PCR
Cell fixation was performed according to the method described by Morohashi and Grotewold (2009) and Morohashi et al. (2012), and chromatin shearing was performed according to the method described by Ohashi-Ito et al. (2014). Cultured cells were fixed for 20 min in fix buffer [0.4 M sucrose, 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 1% protease inhibitor cocktail for plant cell and extracts (Sigma–Aldrich, St. Louis, MO, USA), and 1% formaldehyde]. After addition of 0.1 M glycine and incubation for 5 min with gentle shaking, cells were collected by centrifugation, washed twice with ice-cold water, frozen in liquid nitrogen and stored at −80°C. Frozen cells collected from 2-ml cell cultures were ground using Shake Master neo (Biomedical Science, Tokyo, Japan) at 1,500 rpm for 2 min, and suspended in 1 ml of lysis buffer [50 mM HEPES, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% deoxycholate, 0.1% SDS, 1 mM phenylmethylsulfonyl fluoride, 10 mM sodium butyrate, 1% protease inhibitor cocktail for plant cell and extracts (Sigma-Aldrich)]. Subsequently, the suspension was sonicated for 10 min with Covaris S220 and centrifuged at 15,000 × g for 10 min at 4°C. Immunoprecipitation was performed with 240 µl aliquots of the supernatants and 8 µl of anti-GFP antibody (ab290, Abcam, Cambridge, UK) using OneDay ChIP kit (Diagenode, Liège, Belgium) according to the manufacturer’s protocol. Input samples were prepared from 24 µl aliquots of the supernatants. Real-time quantitative PCR was run using LightCycler480 Probes Master (Roche) with sets of Universal ProbeLibrary probes (Roche) and primers listed in Supplementary Table S1 on a LightCycler480 instrument II (Roche) according to the manufacturer’s protocol.
Quantification of endogenous IAA levels
The culture cells harboring the estrogen-inducible LHW and T5L1-GFP transgene were used for the quantification of endogenous IAA levels. Induced-cells were treated with 5 µM estrogen for 24 h, washed with culture medium three times, and then continued to culture. Quantification was performed according to Kojima et al. (2009).
Histological analysis and fluorescence imaging
GUS staining of 8-day-old seedlings was performed as described previously (Ohashi-Ito et al. 2010). DIC images were taken using a BM5500 microscope (Leica Microsystems, Nussloch, Germany). Technovit sections were prepared using 7-day-old roots according to previously described methods (Ohashi-Ito et al. 2013) and stained with Toluidine Blue O. Sections were prepared from roots at 3 mm above the root tips. Fluorescence images were taken using a FV1200 confocal microscope (Olympus, Tokyo, Japan). For staining the plasma membrane, roots were incubated in 0.01 mg/ml propidium iodide.
Funding
The Ministry of Education, Culture, Sports, Science and Technology of Japan [25113004 to K.O.-I. and 15H05958 to H.F.] and the Japan Society for the Promotion of Science [16H06377 to H.F.].
Acknowledgments
We thank Minobu Shimizu, Yukiko Sugisawa and Sumika Tsuji-Tsukinoki for technical support. We also thank Nam-Hai Chua, Bruno Müller and ABRC for materials.
Disclosures
The authors have no conflicts of interest to declare.
