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Keywords:

  • Arabidospsis thaliana;
  • auxin gradients;
  • cell competence;
  • developmental window;
  • lateral root initiation;
  • Solanum lycopersicum

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
  • Root system architecture depends on lateral root (LR) initiation that takes place in a relatively narrow developmental window (DW). Here, we analyzed the role of auxin gradients established along the parent root in defining this DW for LR initiation.
  • Correlations between auxin distribution and response, and spatiotemporal control of LR initiation were analyzed in Arabidopsis thaliana and tomato (Solanum lycopersicum).
  • In both Arabidopsis and tomato roots, a well defined zone, where auxin content and response are minimal, demarcates the position of a DW for founder cell specification and LR initiation. We show that in the zone of auxin minimum pericycle cells have highest probability to become founder cells and that auxin perception via the TIR1/AFB pathway, and polar auxin transport, are essential for the establishment of this zone.
  • Altogether, this study reveals that the same morphogen-like molecule, auxin, can act simultaneously as a morphogenetic trigger of LR founder cell identity and as a gradient-dependent signal defining positioning of the founder cell specification. This auxin minimum zone might represent an important control mechanism ensuring the LR initiation steadiness and the acropetal LR initiation pattern.

Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

The root system integrates many signals that provide the plant with an advantage to select growth directions in a changeable environment (Malamy, 2005; Laplaze et al., 2007). Root architecture results from a reiterative process of lateral root (LR) formation. As LR organogenesis takes place in an extremely dynamic environment within the simultaneously growing parent roots, an important question is how the acropetal pattern of root initiation is established and maintained and what are the mechanisms controlling recurrent initiation of new LR primordia (LRP).

Auxin is a key regulator of primary (Sabatini et al., 1999; Friml et al., 2002), lateral (Celenza et al., 1995; Laskowski et al., 1995, 2008; Casimiro et al., 2001; Benkováet al., 2003; Dubrovsky et al., 2008), and adventitious (Boerjan et al., 1995; Ullah et al., 2003; Sorin et al., 2005) root development. In Arabidopsis thaliana and most other eudicots, LR formation starts from pericycle cells adjacent to a xylem pole (Laskowski et al., 1995; Dubrovsky et al., 2000; Beeckman et al., 2001). Consistent with the important role of auxin in root development, several mutants in various components of the auxin signaling machinery either lack or have a reduced LR initiation. In the presence of auxin, the transcriptional repressors AUX/IAA interact directly with the auxin receptor, TRANSPORT INHIBITOR RESPONSE1 (TIR1), and are targeted for degradation, thus permitting AUXIN-RESPONSE FACTOR (ARF) proteins to regulate transcription of auxin-responsive genes. Gain-of-function mutations in IAA14/SLR (Fukaki et al., 2002, 2005; Vanneste et al., 2005) and IAA28 (Rogg et al., 2001; Dubrovsky et al., 2009) are deficient or strongly affected in LR initiation, respectively. The double arf7 arf19 mutant does not form any LRs (Okushima et al., 2005; Wilmoth et al., 2005) and is incapable of LR initiation (Okushima et al., 2007; Dubrovsky et al., 2009). Besides transduction, the polar auxin transport represents an important degree of control of the auxin activity. Auxin influx (Bennett et al., 1996; Yang et al., 2006) and efflux (Luschnig et al., 1998; Noh et al., 2001; Petrášek et al., 2006) carriers are main components of the polar auxin transport machinery regulating auxin distribution in tissues and organs, whereas chemical or genetic interference with the polar auxin transport affects LR initiation (Casimiro et al., 2001; Benkováet al., 2003). Although the role of auxin in founder cell specification and LRP initiation is clearly established, the mechanisms regulating the spatiotemporal periodicity of auxin-dependent LRP initiation events are still not elucidated.

Lateral root formation is a complex process that comprises several growth control points. Protoxylem-adjacent pericycle cells in the young differentiation zone of Arabidopsis are capable of proliferating without intervening mitotic quiescence after they leave the root apical meristem (Dubrovsky et al., 2000). This and previous studies (Gladish & Rost, 1993) have suggested that the predetermination of pericycle cells and their subsequent participation in LRP formation are part of a continuous process that starts in the root apical meristem, or soon after cells exit the meristem. Indeed, recent research has suggested that the first growth control point in LR development consists of ‘priming’, that is, determination of pericycle cells for future LRP formation. It takes place close to the root apical meristem, namely in the elongation zone: regular fluctuations in auxin responsiveness in this root zone were found to correlate with subsequent positioning of LRs (De Smet et al., 2007; De Rybel et al., 2010; Moreno-Risueno et al., 2010). Apparently, developmental changes may occur in pericycle cells at this time; nevertheless, it is still unknown how activation of auxin response in the protoxylem cell layer (De Smet et al., 2007) causes the priming in the pericycle cell layer. Interestingly, the auxin responsiveness simultaneously oscillates with activity of two different sets of genes, one in phase and the other in antiphase (Moreno-Risueno et al., 2010).

A second growth control point in the process of root formation consists of specification of LR founder cells and their subsequent divisions, forming of an early-stage primordium. These events occur in the root differentiation zone soon after completion of the cell elongation. Time-lapse analysis in Arabidopsis has shown that LR initiation only happens within a narrow time window (Dubrovsky et al., 2006a). This developmental window (DW) was defined as a period during which pericycle cells in the young differentiation zone remain in a state that allows LR founder cell specification (Dubrovsky et al., 2006a). This specification has been associated with increase in auxin response in small groups of protoxylem-adjacent pericycle cells (Benkováet al., 2003). Live imaging has confirmed that these auxin-responsive pericycle cells are LR founder cells, because soon after they express the auxin-response reporter DR5rev:GFP, these cells divide and form a primordium (Dubrovsky et al., 2008). Recently, the auxin-responsive GATA23 transcription factor has been demonstrated to become active in founder cells c. 10 h after priming of protoxylem cells in the elongation zone; expression of GATA23 depends on the IAA28 function (De Rybel et al., 2010). When a stage I LRP is formed, the third growth control point starts to operate, which regulates LRP morphogenesis and patterning. These processes depend on the AP2/EREBP gene PUCHI (Hirota et al., 2007) and on BDL/IAA12 and MP/ARF5 genes (De Smet et al., 2010).

Previously, we have shown that in the Arabidopsis primary root, LRP initiation occurs in a regular acropetal pattern at the beginning of the differentiation zone and that each new initiation event always takes place distally to a previous one in the direction of the root apex (Dubrovsky et al., 2006a). Here, we wondered how the DW is established and maintained. To achieve a typical acropetal root branching pattern, founder cell specification must be controlled in both time and space. Pericycle cell proliferation is rarely found between developed LRs (Dubrovsky et al., 2000), indicating that pericycle cells located outside the DW seldom participate in the initiation of new LRP. External auxin treatment can induce LRs along the entire parent root, overcoming the acropetal pattern observed during root growth under standard growth conditions (Goldacre, 1959; Blakely et al., 1982; Laskowski et al., 1995). Therefore, we hypothesized that the maintenance of this acropetal pattern must require a still unknown spatiotemporal auxin distribution along the root, spatiotemporal restrictions of the auxin activity, or differential auxin sensitivity along the parent root. To test this hypothesis, we analyzed the endogenous auxin response and auxin content gradients along the parent root, and tested how these gradients correlate with the site of LRP formation. We found that an auxin minimum zone in the primary root corresponded to a zone in which pericycle cells could acquire with the highest probability a founder cell identity. When the gradients in the auxin distribution or perception along the root were altered genetically or pharmacologically, regular acropetal pattern of LRP formation could no longer be maintained.

Materials and Methods

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Transgenic lines, growth conditions and treatments

Transgenic Arabidopsis thaliana (L.) Heynh. lines in Columbia-0 (Col-0) background were CYCB1;1DB::GUS (Colón-Carmona et al., 1999), DR5::GUS (Ulmasov et al., 1997), DR5rev::GFP (Friml et al., 2003), DR5rev::GFP in tir1 afb2 afb3 triple mutant background (Dharmasiri et al., 2005b), and pIAA14::mIAA14-GR (Fukaki et al., 2005). Tomato (Solanum lycopersicum L.) plants cv Ailsa Craig were used. To generate a tomato pIAA2::GUS line, seedlings were transformed with the p1G4 construct carrying the Arabidopsis pIAA2::GUS reporter (a kind gift from J. Normanly, University of Massachusetts, MA, USA). The DR5::GUS tomato line had been reported previously (Dubrovsky et al., 2008).

Unless otherwise indicated, plants were grown in vertically oriented Petri dishes on 0.2 × MS (Murashige & Skoog, 1962) medium, pH 5.7, supplemented with 1% (w/v) sucrose (Dubrovsky et al., 2006a). To analyze LR initiation following auxin transport inhibition, Arabidopsis DR5::GUS seedlings were grown on 10 μM 1-naphthylphthalamic acid (NPA) for 10 d, the 1.5 mm apical and basal root segments were removed, and the remaining root divided into three equal portions, which were transferred onto MS medium with or without auxins for three additional days. Tomato DR5::GUS seedlings were grown in liquid MS medium with 10 μM NPA for 6–7 d, shoots and root meristems were removed, and roots transferred on to solid control medium for 3 d. Roots were cleared as described (Dubrovsky et al., 2006a, 2009).

Dexamethasone (DEX)-inducible transient inhibition of LR initiation was tested in segmented-agar dishes with 50 μM DEX added only to the lower half of the vertically oriented dish (Supporting Information, Fig. S3a). Six-day-old pIAA14::mIAA14-GR seedlings were transferred on the segmented medium so that 1 mm of the root tip was on the (+) DEX sector. Root portions grown on a DEX sector for 3 d were analyzed either immediately or after transfer on to MS medium with or without naphthalene-acetic acid (NAA) for two additional days.

To estimate the DW duration for LR initiation, 24 h root growth increments between days 7 and 8 were measured and the average rate of root growth calculated as (mm h−1). Roots were cleared and the distance (d) was measured from the quiescent center (QC) to the most distal LRP. The DW duration (h) was estimated as = (dmax − dmin)V−1, where dmax and dmin (mm) are the maximal and minimal distances, respectively.

Quantification of DR5rev::GFP expression along the Arabidopsis root and microscopy

DR5rev::GFP expression was analyzed by confocal laser scanning microscopy (CLSM). Live roots were analyzed with or without staining with 4 μM neutral red at pH 5.7 (Dubrovsky et al., 2006b); some roots were fixed for 6–12 h at 4°C in 3.7% para-formaldehyde in phosphate-buffered saline (pH 7.4), supplemented with 1 μg ml−1 propidium iodide. Confocal images spanning the live root from the tip to the emerged LRs were acquired with a 10× objective under the same settings. Confocal sections of 60 μm were taken, corresponding to the diameter of the root central cylinder. Images were assembled in Adobe Photoshop (Adobe Systems, San Jose, CA, USA). Green fluorescent protein (GFP) intensity was measured with ImageJ (http://rsb.info.nih.gov/) as pixel density within the tissues of the central cylinder (except pericycle) starting from the QC and the background level was subtracted. Pixel densities within the QC cells were considered at 0 μm. For the most distal 400 μm of the root, intensities were measured in portions of 0–100, 100–200, and 200–400 μm and averaged over three measurements per portion. Intensities of three consecutive 167 μm portions were measured for each 500 μm interval. Values were expressed as percentage of mean pixel density relative to the GFP maximum.

Microscopy and image acquisition were done as described (Dubrovsky et al., 2006b). Distances from the QC to the most distal primordium were measured on cleared roots prepared as previously described (Dubrovsky et al., 2009) with an ocular micrometer. Founder cells and most distal primordia were detected by accumulation of the DR5rev::GFP activity (Dubrovsky et al., 2008) under a CLSM with a 40× or a 63× objective. Distances from the QC to the founder cells and the most distal primordia detected were measured under CLSM with a 10× objective and LSM5 Image Examiner program (Zeiss) to the precision of 1 μm.

Analysis of indole-3-acetic acid content and transport

Endogenous free IAA was measured after methylation and GC-MS analysis as previously described (Kim et al., 2007) in three independent experiments of 10 seedlings each. The auxin transport assay was done as previously described (Geisler et al., 2005; Peer & Murphy, 2007), except that the Col-0 seedlings were initially grown for 5 d under 120 μmol m−2 s−1 light in Petri dishes with 0.25 × MS, supplemented with 0.5% sucrose at pH 5.5. Before the assay, seedlings were transferred to 2-mm-wide, vertically discontinuous filter paper strips saturated with liquid 0.25 × MS medium and were allowed to equilibrate for 2 h under yellow light. With a micromanipulator, a 10 nl microdroplet containing 1 μM unlabeled IAA and 1 μM [3H]IAA (specific activity 20 Ci mmol−1, American Radiochemical, St Louis, MO, USA) in dimethylsulfoxide was applied on the shoot apex. Seedlings were incubated in yellow light for 4, 4.5, 5, 6 and 7 h, after which the hypocotyls and cotyledons were removed, and 2 mm root sections sequentially harvested and assayed by scintillation counting (Tri-Carb 3180; Perkin Elmer, Waltham, MA, USA). A 5 h time point was chosen for the measurements of transported auxin because, under the conditions used, auxin could be detected first at the root apex (> 2 × background) at 4.5 h. Samples were collected separately, extracted with 100% metanol, and analyzed according to Östin et al. (1998) for IAA, 2-oxoindole-3-acetic acid (IAAox), and 2-oxoindole-3-acetyl-β-d-O-glucopyranose (IAAox-Gluc).

pIAA2::GUS expression in tomato root

Roots of 6-d-old pIAA2::GUS seedlings were fixed in 50 mM phosphate buffer, pH 7.2, containing 0.3% formaldehyde and 0.3 M mannitol for 10 min, washed three times with 50 mM phosphate buffer, and incubated overnight in 100 mM phosphate buffer (pH 7.2), 0.5 mM K3[Fe(CN6)], 0.5 mM K4[Fe(CN6)], 10 mM EDTA, 20% methanol, 0.01% Triton X100, and 2 mg ml−1 X-gluc (Gold BioTechnologies, St Louis, MO, USA). After clearing, roots were mounted in 25% glycerol and analyzed.

For fluorometric GUS expression assay, 1 mm fragments of 10 roots were cut at positions as indicated in Fig. 1(e) with a dissecting microscope, collected on ice, and ground in 100 μl extraction buffer containing 40 mM phosphate buffer, pH 7.2, 1 mM EDTA, 0.01% Triton X100, and 0.01% sodium lauroyl sarcosine. Samples were centrifuged at 10 000 g for 10 min, and 10 μl from each sample was mixed in duplicates with 40 μl reaction buffer containing 1 mM dithiothreitol and 3 mg ml−1 4-methylumbelliferyl-β-d-glucuronide (Marker Gene Technologies, Eugene, OR, USA). Reactions were incubated at 37°C for 20 min, cooled to room temperature, and stopped by 160 μl 0.2 M Na2CO3. Fluorescence was measured with a Synergy 2 Multi-Detection Microplate Reader (Winooski, VT, USA) and excitation and emission at 360 and 465 nm, respectively. Protein concentration was measured with a Micro BCA Protein Assay Kit (Thermo Scientific, Rockford, IL, USA) according to the manufacturer’s instructions. GUS expression was calculated as relative fluorescence units mg–1 protein. Values were averaged from two independent experiments.

image

Figure 1. Gradients in auxin response along Arabidopsis and tomato roots. (a) Gradient of DR5rev::GFP activity in 7-d-old Arabidopsis root; numbers indicate distance from the quiescent center (QC) (mm). (b, c) DR5 activity measured in the central cylinder as relative mean pixel density of the green fluorescent protein (GFP) signal and expressed as a percentage of maximum activity (measured in the QC (for b)) and in the proximal part of the root (for c) of each individual root in 5-d-old (= 7) (b) and 14-d-old (c) Arabidopsis plants (n = 5). Insets show the mean pixel density within the first 400 μm from the QC. (d, e) Gradient in the pIAA2::GUS auxin reporter activity along the primary root of tomato. (d) pIAA2::GUS reporter expression in a 6-d-old seedling; numbers indicate distance from the QC (mm). (e) Quantification of GUS expression with a fluorometric assay in 1 mm root segments collected at the indicated distances. Values are averages from two independent experiments with 10 roots each. Mean ± SEs are shown (b, c, e).

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Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

LR founder cell specification is restricted to a narrow developmental zone in the parent root

Previously, we have shown that, in Col-0 plants, LR initiation occurs within a narrow root portion behind the root tip and have calculated that each subsequent LRP is initiated within a time-frame interval of c. 10 h, referred to as the ‘developmental window for LR initiation’ (Dubrovsky et al., 2006a). Spatially, this time-frame should correspond to a specific zone a few mm above the QC. In the Col-0 wild-type, the distance from the QC to the most distal LRP ranged from 2.4 to 6.8 mm, defining a zone with the highest capability for LRP initiation, corresponding to the DW (Fig. 2). In terms of time this DW corresponds to 8.5 h. Because the earliest known sign of LR founder cell identity is the increased DR5 activity in pericycle cells (Benkováet al., 2003; Dubrovsky et al., 2008), we used DR5rev::GFP to detect founder cells by CLSM and established their position relative to the most distal primordium in the same roots (Table 1). The distance between the site of founder cell specification and that of primordium initiation was short, on average 0.13–1.1 mm, indicating that once the founder cell specification occurs, primordium initiation follows relatively soon after. These data show that founder cell specification and primordium initiation in Arabidopsis (Col-0) occur, on average, at 4.21 ± 1.01 mm (mean ± SD, n = 77, 8-d-old plants) from the QC, i.e. in the young differentiation zone. Thus, both of these processes are separated in time and space from the pericycle cell priming that takes place within the elongation zone (De Smet et al., 2007; Moreno-Risueno et al., 2010), that is, at c. 0.2–1.2 mm from the QC in Arabidopsis (Tapia-López et al., 2008; J. G. Dubrovsky, pers. obs.). Thus, founder cell specification and LRP initiation represent a growth control point in the LR formation that operates in the differentiation zone.

image

Figure 2. Developmental window operating during lateral root initiation shown as the distribution of distances from the quiescent center (QC) to a most distal primordium in 8-d-old Arabidopsis thaliana Col-0 plants (= 77).

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Table 1.   Location of founder cells and distal lateral root primordium detected along the root of Arabidopsis DR5rev::GFP line
Plant ageDistance to founder cells (mm)Distance to distal lateral root primordium (mm)
MinimalMaximalMeanMinimalMaximalMean
  1. Data are means ± SE, n is indicated in the parentheses. The distance was measured from the quiescent center (QC). In 14-d-old plants, founder cells were slightly more distant from the QC than in 7-d-old plants (= 0.007, Student’s t-test); the distance to the distal lateral root primordium was the same in 7 and 14-d-old plants (= 0.753, Student’s t-test).

7 d2.7244.8613.348 ± 0.198 (12)3.3365.4024.404 ± 0.260 (9)
14 d2.9205.1124.170 ± 0.198 (14)3.0105.2704.299 ± 0.204 (14)

Auxin reporters reveal an auxin response minimum along the primary root

Because exogenous auxin can activate pericycle cells and promote new LR formation along the entire parent root, whereas in the intact root initiation can only take place in a narrow zone, we wondered whether endogenous auxin distribution and response can be related to the establishment of the DW for founder cell specification. In the Arabidopsis root, two auxin maxima have been reported, one in the QC (Sabatini et al., 1999; Friml et al., 2002; Ljung et al., 2005; Petersson et al., 2009) and another at the root base (Bhalerao et al., 2002; Marchant et al., 2002; Ljung et al., 2005), forming distal and proximal auxin gradients, respectively. As not much is known about the auxin response distribution in the root zones where LRP initiate and develop, the expression of the auxin-sensitive DR5rev::GFP reporter (Friml et al., 2003) was analyzed along the entire root of Arabidopsis with CLSM (Fig. 1a). We observed the DR5 maximum in the QC cells as previously reported (Sabatini et al., 1999). Within the root meristem, the DR5 activity was maintained in a one-cell-thick layer of immature xylem-precursor cells, occurred at a low level in elongating and differentiating protoxylem cells, and was not detected in pericycle cells (Fig. S1). Starting at c. 1.2 mm from the root tip, no DR5 activity was detected in the young differentiation zone (Fig. S1). Quantification of DR5rev::GFP expression confirmed that its level became very low at 0.2 mm from the QC (insets in Fig. 1b,c) and that, at 5–6 mm from the root tip, the DR5 activity started to steadily increase in a proximal direction within the tissues of the root central cylinder, excluding pericycle cell layer (Fig. 1a–c). This gradient seems to be independent of seedling age, because the DR5 activity had a similar pattern in roots of 5-d-old (Fig. 1b), 7-d-old (data not shown), and 14-d-old (Fig. 1c) seedlings. A similar pattern of auxin response was also observed based on fluorimetric GUS measurements in roots of tomato expressing the pIAA2::GUS auxin reporter (Fig. 1d,e). Thus, there is a well defined auxin response minimum zone that corresponds spatially to the DW for LR initiation that is found at the start of the differentiation zone in both Arabidopsis and tomato.

Monitoring auxin content and distribution along the root

Free IAA concentrations were assayed in primary roots of 5-d-old Col-0 seedlings. Steady-state free IAA concentrations decreased progressively in the primary root from the shoot-to-root transition zone downward to the root apex and then increased again at the root apex (Fig. 3). The lowest free IAA content was detected in root segments taken 2–4 mm from the apex, which was 40% less than that in the 12–14 mm root segments (= 0.002, Student’s t-test), and was accompanied with a concurrent increase in the oxidative breakdown products IAAox and IAAox-Gluc (data not shown). This pattern of free auxin distribution closely corresponded to the pattern we detected in DR5 and IAA2 promoter activities along the root of Arabidopsis and tomato, respectively (Fig. 1), and is in agreement with previous reports on overall auxin distribution in the root (Ljung et al., 2005; Petersson et al., 2009).

image

Figure 3. Transport of shoot-derived auxin along the root and free auxin content. The line plot represents radioactively detected signal in pooled 2 mm segments from 10 seedlings (= 6) 5 h after 10 fmol 3[H]IAA was deposited at the shoot apex of 5.5-d-old seedlings. The asterisks indicate differences in each subsequent root segment when compared with the previous segment at < 0.005 (Student’s t-test). The histogram represents free IAA content of seedlings sampled in the same manner after treatment with a solvent control (= 3). The asterisk on the histogram indicates a difference between the root segments of 0–2 and 2–4 mm (= 0.001, Student’s t-test). Mean ± SEs are shown.

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To correlate the steady-state free auxin concentrations with the auxin transport activity, we performed nanoscale polar auxin transport assays from the shoot to root apex. In these assays, 10 nl [3H]IAA (1 μM, 20 Ci mmol−1) was applied to the apex of 10 seedlings that were incubated under dim yellow light. Starting at 2 h, 2 mm sections were excised serially from the root apex upward to the root-to-shoot transition zone. The [3H]IAA could first be detected (by scintillation counting of the pooled sections) at the primary root apex (> 2 × background concentrations) at 4.5 h and signals increased progressively in the pooled sections closer to the root-to-shoot transition zone. However, at 5 h and later, [3H]IAA signals at the root apex increased steadily and decreased progressively in the rest of the root (data not shown) in a manner consistent with the observed steady-state free IAA concentrations (Fig. 3). Altogether, the auxin-reporter analysis and the direct measurements of auxin content and transport reveal that auxin concentrations and responses along the root obey a certain regularity that is probably maintained through polar auxin transport. An auxin minimum is defined between a distal auxin gradient at the root tip, and a proximal gradient along the central cylinder in Arabidopsis and tomato roots.

The zone of auxin minimum in the root overlaps with the developmental window for founder cell specification and primordium initiation

As already described, the distance from the most distal primordium to the QC was between 2.4 and 6.8 mm in Arabidopsis (Fig. 2) in a zone overlapping that of minimal auxin response and content (between 1.5 and 5–6 mm from the QC), from which the proximal auxin gradient starts (Table 1; Figs 1, 3). Similarly, in tomato roots, the distance from the QC to the most distal LRP ranged from 4.9 to 10.7 mm (mean ± SE, 7.9 ± 0.30 mm, = 23), corresponding to a zone with low auxin response (Fig. 1d,e). When we plotted the distance from the QC to the most distal LRP against the distance to the start of the proximal auxin gradient for each root, a clear correlation was found for both Arabidopsis DR5rev::GFP plants (= 0.80; Fig. 4a) and tomato pIAA2::GUS plants (= 0.83; Fig. 4b). Thus, founder cell specification and initiation of an early-stage primordium correlates statistically with the start of the proximal auxin gradient.

image

Figure 4. Correlation between the lateral root (LR) initiation site and the start of the proximal auxin gradient in Arabidopsis and tomato root. (a) Correlation between the start of the proximal auxin gradient (expressed as 5% of maximum auxin response in the quiescent center (QC)) and the position of the most distal LR primordia (LRP) in 7-d-old DR5rev::GFP Arabidopsis plants and pin3 and pin7 mutants crossed with DR5rev::GFP. (b) Correlation between the start of the detectable pIAA2::GUS expression and the position of the most distal LRP in 6-d-old tomato plants. Regression line and correlation coefficient are shown (= 7–8 (a) and = 23 (b)). For Arabidopsis and tomato wild-type, < 0.05 (Pearson’s correlation test); for pin3 and pin7, P = 0.273 and 0.908, respectively (Pearson’s correlation test).

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Lack of PIN3 and PIN7 auxin efflux carriers has been demonstrated to interfere with LR initiation (Benkováet al., 2003). As both PIN3 and PIN7 participate in the regulation of the polar auxin transport (Friml et al., 2002, 2003), we examined whether the zone of auxin minimum in pin3 and pin7 mutants was altered compared with that of the wild-type and, if so, how this correlated with the positioning of the LR initiation site. In both pin3 and pin7, the DR5rev::GFP signal behind the root elongation zone started at a shorter distance from the QC, namely 2–3 and 3–4 mm from the QC, respectively, while it was between 4 and 7 mm from QC in the wild-type (Fig. 4a). In pin3 and pin7, there was low (= 0.44) or no correlation (= 0.04), respectively, between relative positions of the most distal LR initiation events and the start of the proximal auxin-response gradient (Fig. 4a). In both of these cases, the correlation was not statistically significant (> 0.05, Pearson correlation test) in contrast to the wild-type Arabidopsis and tomato seedlings (< 0.05, Pearson correlation test), indicating that in the wild-type, maintenance of the auxin minimum zone in the root central cylinder depends on the polar auxin transport and that the auxin minimum zone defines the DW for the LR initiation. When the zone becomes altered, the correlation between the sites of primordium initiation and location of the auxin minimum zone is lost.

Pericycle cells have the highest probability to become founder cells in the auxin minimum zone

If the zone of the auxin minimum defined the DW, we hypothesized that pericycle cells transiently present in this zone would be able to acquire founder cell identity and initiate a LR, and that once they had moved out of the zone as a result of the primary root growth, this ability would disappear under normal growth conditions. To test these hypotheses, we transiently inhibited the LRP initiation and examined the pattern of its restoration after removal of the restrictive conditions. We first used NPA to block the LRP initiation (Casimiro et al., 2001; Himanen et al., 2002, 2004) during the first 6 d of root growth. When the seedlings were transferred onto NPA-free medium, initiation was restored exclusively in the root portion formed after the inhibitor removal (Fig. 5a), suggesting that a new DW with pericycle cells competent for LR initiation had been established at the root tip. The IAA polar transport was restored within 24 h after NPA removal (Fig. S2), thus confirming the transient character of the NPA inhibition of the polar auxin transport. After transfer to NPA-free medium, no LRP were detected in the zone grown in the presence of NPA, even after prolonged incubation without the inhibitor, demonstrating the irreversible loss of the capability to initiate a primordium in the root portion where the polar auxin transport had been inhibited. Next, founder cell activation was inhibited in a genetically controlled manner with the pIAA14::mIAA14-GR Arabidopsis line (Fukaki et al., 2005) in which a stabilized form of the IAA14/SLR repressor protein accumulates in the nucleus upon DEX treatment (Fukaki et al., 2005). LR formation in the pIAA14::mIAA14-GR seedlings was inhibited transiently upon the DEX treatment (De Smet et al., 2007), but it is unclear whether LRP initiation was abolished. In roots grown on agar plates with DEX-supplemented sectors (Fig. S3a), LRP initiation was not completely eliminated in the presence of DEX (Fig. S3b,c), but the density of the pericycle activation events (including LR and LRP) was reduced 1.8-fold in the root portion growing on the sector with 50 μM DEX (Fig. 5b). Importantly, when seedlings were transferred to DEX-free medium for an extra 2 d, the density of pericycle activation events did not increase (> 0.05, Student’s t-test) in the DEX-treated root portion, confirming that the loss of ability to initiate LRP is irreversible once pericycle cells exit the auxin minimum zone. However, the density of pericycle activation events in the DEX-treated segment was increased 12.9-fold upon seedling transfer to NAA-containing medium (Fig. 5c), demonstrating that the competence for founder cell specification was not lost. Thus, these data provide consistent experimental evidence that LR initiation is restricted to a DW. Although cells outside the DW retain their competence to acquire founder cell identity, they require additional auxin stimulation.

image

Figure 5. Irreversible nature of the transient inhibition of the lateral root (LR) initiation in Arabidopsis. (a) Col-0 seeds germinated and seedlings grown on medium supplemented with 10 μM 1-naphthylphthalamic acid (NPA) for 6 d; seedlings were transferred to NPA-free Murashige–Skoog (MS) medium for an additional 3 d; LRs and primordia were formed exclusively in the root portion formed after transfer. The numbers of both primordia and lateral roots present within the root portions formed before and after treatment are shown in one of three independent experiments (= 22). (b) Six-day-old pIAA14::mIAA14-GR were transferred on to plates either without (−) or with 50 μM dexamethasone (DEX) (+) sectors for 3 d. Seedlings had decreased density of pericycle activation events (LRs and primordia) in the root portion formed on DEX-supplemented (+) sectors. (c) After DEX treatment, seedlings were transferred on to DEX-free medium without (−) or with 1 μM (+) naphthalene-acetic acid (NAA) for an additional 2 d (see Supporting Information, Fig. S3, for experimental setup). On NAA-free medium, the density of pericycle activation events did not increase in the previously DEX-treated root portion, but it increased upon transfer to NAA-containing medium. Mean ± SE (= 19–23) of two independent experiments. (+) or (−) DEX treatments differed at < 0.001 (Student’s t-test).

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Indeed, a temporal 9 h application of Sephadex beads loaded with 10 μM NAA at different portions of the root further confirmed that the highest number of initiation events occurred in the most distal (1–10 mm) root zone, comprising the zone of auxin minimum (Fig. S4), and dramatically decreased with distance from the root tip. Dynamics of pericycle activation of NPA-pretreated roots induced by NAA revealed that pericycle cells were activated first, within 2 h of the treatment, in the root portion corresponding to the DW (within the apical 5 mm of the root). However, a 6 h auxin treatment was necessary before LR initiation events appeared in the more proximal parts of the primary root (Fig. S5). These experiments demonstrate that pericycle cells in the auxin minimum zone and within the DW portion have the highest auxin responsiveness compared with other parts of the root. Owing to the Casparian strip in the endodermis cells, we could not exclude that the pericycle was less accessible to NAA in mature root portions than in the root tip. To facilitate auxin access to the pericycle, we also analyzed auxin responsiveness in three physically separated consecutive root segments lacking LRPs after NPA treatment. In both untreated and auxin-treated root segments, the density of pericycle activation events was the greatest in the apical root segment (with the meristem and elongation zone excised), comprising the auxin minimum zone, whereas in the middle and basal segments, initiation progressively decreased (Fig. 6). A similar gradual decrease in induced initiation events outside the DW was found in tomato DR5::GUS seedling roots (Fig. S6). This analysis reveals that although all pericycle cells along the root of young plant seedlings can respond to auxin with pericycle activation, the probability of pericycle cells to acquire a founder cell identity is the greatest in the root portion corresponding to the zone of auxin minimum. Hence, in the auxin minimum zone, the developmental state of the pericycle cells makes them more sensitive to external auxin and the enhanced pericycle activation in this zone depends on the original auxin status of this tissue. Overall, our data show that under normal growth conditions the ability to acquire a founder cell identity is transient in the auxin minimum zone and lost when the cells exit this zone. When auxin is provided externally, the pericycle is activated both inside and outside the DW, but the cells in the zone of auxin minimum are still more prone to LR formation.

image

Figure 6. Pericycle cell competence to form lateral roots (LRs). DR5::GUS Arabidopsis seedlings were grown on 10 μM 1-naphthylphthalamic acid (NPA) for 10 d, and then 1.5 mm segments at the root base and at the root apex were excised and discarded. Then the root was divided into three approximately equal root segments: basal, median, and apical. Every cut is marked with an X on the scheme. The root segments were transferred for 3 d on to control (= 29–33; white bars), 0.1 μM 2,4-Dichlorophenoxyacetic acid (= 22–33; gray bars), or 1 μM naphthalene-acetic acid (NAA) (= 20–28; black bars) media; the density of pericycle activation events (LRs and primordia) per mm of the root is shown. The combined data of three independent experiments are shown. Means ± SE are shown. Different letters indicate the difference between apical, median, and basal segments within the same treatment at < 0.05 (Kruskal–Wallis one-way ANOVA on ranks). The average length of the root segments was 4.8 ± 1.2 mm (mean ± SD; = 146).

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Auxin perception is required for proper spatial pattern of primordium initiation

TRANSPORT INHIBITOR RESPONSE1 (TIR1), and the homologous auxin receptors AUXIN-SIGNALING-F-BOX1 (AFB1), AFB2 and AFB3 regulate developmental responses to auxin (Dharmasiri et al., 2005a,b; Kepinski & Leyser, 2005). To examine their impact on LR initiation, we studied the tir1 afb2 afb3 mutant in the DR5rev::GFP background and found, in agreement with published results (Parry et al., 2009), that in 70% of the seedlings the primary root was shorter than 3 mm (= 96). Therefore, we analyzed only those mutant seedlings that had an elongated primary root, on average 22.1 mm (Table 2). As DR5 activity is not detected in this triple mutant (Dharmasiri et al., 2005b), we could not use this reporter to detect founder cells and to monitor their position relative to the auxin gradients along the primary root. However, we observed that the mutant seedlings exhibited an irregular primordium initiation that occurred at an increased distance from the QC and varied among the seedlings (Table 2). In other words, the DW for founder cell specification was apparently less precisely defined in the mutant than in the wild-type. We also found various defects in pericycle cell development and primordium morphology, including pericycle cell division without LRP formation (Fig. 7d), two-cell layered pericycle (Fig. 7e), and abnormal LRP formed from two-cell layered pericycle (Fig. 7f). Exogenous auxin induced cell proliferation in the xylem-adjacent pericycle cells of the tir1 afb2 afb3 mutant with density of activation events, dynamics, and distribution along the root axis similar to those of untreated roots (Fig. 7g–i). Overall, these data suggest that accurate auxin perception is required for correct establishment of the zone of auxin minimum, DW, and normal LRP initiation process.

Table 2.   Primary root length and lateral root (LR) development in 8-d-old Arabidopsis wild-type (DR5rev::GFP) and triple tir1 afb2 afb3 mutant (DR5rev::GFP)
 Distance fromthe root tip to the distal LRP (mm) (10)Number of plants with emerged LRs (15)Root length (mm) (10)
  1. LRP, lateral root primordia.

  2. Mean ± SD, n is indicated in parentheses in the column headers.

DR5rev::GFP4.87 ± 0.7715/1543.0 ± 4.8
tir1 afb2 afb3 DR5rev::GFP7.36 ± 2.963/1522.1 ± 5.2
Student’s t-test< 0.05 < 0.001
image

Figure 7. Defects in lateral root (LR) initiation and primordium organogenesis in auxin receptor tir1 afb2 afb3 triple mutant in comparison with wild-type (DR5rev::GFP) in 8-d-old Arabidopsis seedlings. (a–c) Wild-type without treatment (= 10). (a) Protoxylem-adjacent pericycle cell. (b) Stage I primordium. (c) Emerging primordium; asterisks mark flanking pericycle cells. (d–f) Defects in untreated roots of the tir1 afb2 afb3 (DR5rev::GFP) line (= 10). (d) Unusually short pericycle cells. (e) Two-layered pericycle found in some root portions; p, epicycle. (f) Abnormal primordium formed from a two-layered pericycle. Asterisks mark the margins of the primordium. (g) Quantitative analysis of pericycle activation events in wild-type (DR5rev::GFP; circles) and in tir1 afb2 afb3 (DR5rev::GFP; triangles) upon 10 μM naphthalene-acetic acid (NAA) treatment. Means and 95% confidence intervals are given (= 10). (h) DR5rev::GFP treated with 10 μM NAA during a 24 h period (= 10). (i) Upon a 6 h treatment with 10 μM NAA, pericycle cells in the tir1 afb 2 afb 3 (DR5rev::GFP) triple mutant become shorter as a result of cell division; at 12 h, a two-layered pericycle or stage-II primordia are formed. At 24 h, division of pericycle cells in the area between primordia or LRs (24 h-1), formation of an abnormal primordia from activation of a two-layered pericycle (asterisk, 24 h-2), and formation of a multilayered pericycle in the distal root portion (24 h-3) are observed. The vertical and horizontal arrowheads indicate end walls of pericycle cells and new cell walls, resulting from periclinal cell division, respectively (= 10). Bar, 50 μm.

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

The developmental importance of the auxin distribution has been demonstrated in different organogenic events (Benkováet al., 2009). Auxin acts as a morphogenetic trigger of LR initiation and organogenesis (Dubrovsky et al., 2008). Here, we demonstrate that auxin gradients along the root control the positioning of the site of founder cell specification and their posterior activation, resulting in the acropetal pattern of the LR formation.

The auxin minimum zone defines the DW for LR initiation

Our detailed analyses of the DR5 activity along the primary root supported by direct auxin measurements revealed that in roots the distal and the proximal auxin gradients are separated by a zone of auxin minimum. The distal gradient in the Arabidopsis root typically ceases at 0.2 mm from the root tip, with the DR5rev::GFP expression turning out to be very low (Fig. 1b,c), and at c. 1.2 mm from the QC, no DR5 activity is detected (Fig. S1). The proximal auxin gradient starts in wild-type Arabidopsis seedlings 4.5 to 6 mm from the QC. A similar pattern was found in tomato roots (Fig. 1d,e). In both species, the zone of auxin minimum overlaps with a DW in which specification of founder cells followed by their division occurs with the highest probability (Table 1; Figs 1, 4).

In NPA-pretreated Arabidopsis roots lacking LRP, new initiation events occur exclusively in the root portion formed after removal of NPA and no LRP formation takes place in the pericycle previously exposed to NPA (Fig. 5a). Absence of new pericycle activation events in the NPA-treated zone is not a consequence of NPA retention in this root portion because the IAA polar transport is restored within 24 h upon elimination of the inhibitor. Accordingly, NPA becomes hydrolyzed and inactivated within 24 h at the root tip of treated seedlings of Arabidopsis (Murphy & Taiz, 1999). Interestingly, in seedlings of Arabidopsis (Fig. 6) and tomato (Fig. S6) that were able to form LRs after NPA treatment, the primary root was separated from the shoot and the meristem was removed. This indicates that endogenous auxin gradients established in intact roots are essential for the irreversible arrest of pericycle cell activation for LRP formation. When these endogenous auxin gradients are broken by excisions, the pericycle acquires the capability for founder cell specification and LRP formation.

Mutants in the PIN3 and PIN7 genes coding for auxin efflux carriers of the PIN family (Friml et al., 2003, 2004) have previously been shown to be defective in LR initiation (Benkováet al., 2003; Dubrovsky et al., 2009). The findings reported here reveal that PIN3 and PIN7 are important players in the maintenance of the proximal auxin gradient and in correct positioning and operation of the DW for LR initiation. In both pin3 and pin7 mutants, the auxin minimum zone is reduced and correlation between the start of the proximal auxin gradient and the site of primordium initiation is absent (Fig. 4a), whereas positioning of the LR initiation is more random.

Temporal accumulation of the mIAA14 repressor strongly reduces the density of pericycle activation events and, as with the transient inhibition by NPA, no additional activation takes place in the DEX-treated root zone at a later time, as would be expected if the acropetal pattern of the LR initiation were not maintained. Accumulation of the mIAA14 protein is not sufficient to prevent the LR initiation when seedlings are treated with NAA (Fig. 5c).

Altogether these findings reveal the existence of a mechanism implicated in the regulation of an acropetal pattern of LRP formation. This mechanism depends on auxin transport and distribution, particularly on a zone of auxin minimum that defines a DW for founder cell specification and LR initiation.

The auxin minimum defines the positioning of the LR initiation

Oscillation in DR5 activity within the elongation zone (De Smet et al., 2007; De Rybel et al., 2010; Moreno-Risueno et al., 2010) and subsequent activation of pericycle cells taking place in the differentiation zone are two key growth control points in the LR formation. The aim of this work was to analyze how the processes of the second growth control point, the founder cell specification and their first divisions, are regulated in space and time. We show that a zone of auxin minimum and a DW defined by this minimum are essential. When the DW was pharmacologically or genetically modified, cells could pass through this part of the root without primordium initiation. Our data demonstrate that at time intervals ≤ 8.5 h, the next pericycle activation event takes place within the zone of auxin minimum and that the cells are triggered to become founder cells. This estimation is close to the time intervals between consecutive oscillation events in gene expression activity at pre-branch sites (Moreno-Risueno et al., 2010). This time-related mechanism seems to be well coordinated with the rate of root growth, and the start of the proximal auxin gradient is apparently displaced towards the root tip at time intervals equal to those between subsequent primordium initiation events.

Our work reveals that, paradoxically, LRP initiation triggered by auxin (Dubrovsky et al., 2008) occurs with the highest probability in a zone of auxin minimum at the tip of the primary root. The possibility that pericycle cells in the DW exhibiting an auxin minimum are preferentially activated for LR initiation might be that reduced auxin concentrations in this zone sensitize pericycle cells to small fluctuations in auxin concentrations. In support of this explanation are the experiments with the tir1 afb2 afb3 mutant that suggest a correct auxin perception is essential for DW establishment. Interestingly, when TIR1/AFB-dependent signaling is inhibited pharmacologically, the newly grown root portion no longer shows GATA23 promoter activity typical for founder cells (De Rybel et al., 2010). Developmental mechanisms involved in founder cell specification and stage I LRP formation are not completely understood. We provide a mechanistic explanation for the link between auxin gradients and pericycle activation events and suggest that an auxin minimum zone restricts the occurrence of founder cell specification and LR initiation in time and space. The establishment of this zone might represent an important control mechanism to ensure steadiness and acropetal pattern of primordium initiation.

A developmental window model for LR initiation defined by auxin gradients established along the parent root

We propose the following model of auxin-dependent establishment of a DW for founder cell specification and primordium formation (Fig. 8). In an intact growing root, two distal and one proximal auxin gradients are established, defining a morphogenetic zone that corresponds to a DW for LR initiation and includes founder cell specification and their first divisions leading to a stage I primordium. Within this zone, a minimum auxin concentration coincides with the maximum probability of cells to become founder cells. This window, between c. 2 and 6 mm from the QC, is dynamic and displaced in the same direction and at the same rate as the root growth rate. NPA-sensitive polar auxin transport and auxin receptors of the TIR1/AFB family are important for this DW operation. When external auxin is applied, pericycle cells outside the DW can be activated because they maintain their competence to become founder cells. Under nonstandard growing conditions, such as exogenous auxin or root damage in the natural environment, the DW is no longer operational and initiation can occur outside the DW zone.

image

Figure 8. Model of developmental window (DW) operation. In an intact growing root, two distal and one proximal auxin gradients are established that define a DW for lateral root (LR) initiation. Within this morphogenetic zone, the minimal auxin concentration correlates with the highest probability of pericycle cells to acquire founder cell (FC) identity. This morphogenetic zone, between 2 and 6 mm from the tip, is dynamic and is displaced in the same direction and at the same rate as the root growth rate. Auxin transport and receptors of the TIR1 family are required for operation of this DW. When external auxin is applied, its distribution is rearranged and cells outside the DW can be activated, because they maintain their competence to form FCs. NPA, 1-naphthylphthalamic acid (NPA).

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In conclusion, our data indicate that a DW in which the LR initiation takes place in an acropetal sequence correlates spatially and temporally with the establishment and maintenance of the auxin minimum zone, which is defined as a zone between distal and proximal auxin gradients and is functionally important to maintain steady LRP formation. Functionality of this window relies on active auxin transport and perception and on response mechanisms.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

We thank J. Murfett for the DR5::GUS line, M. Estelle for the tir1 afb2 afb3 mutant, H. Fukaki for pIAA14::mIAA14-GR, P. Doerner for the CycB1;1DB::GUS line, J. Normanly for the p1G4 AtIAA2::GUS reporter construct, A. Saralegui, A.L. Martínez-Valle and S. Ainsworth for excellent technical help, N. Doktor for help with artwork and pixel-density analysis, P. Benfey for critical reading of the manuscript, and M. De Cock for help in preparing it. This research was supported by the European Research Council starting independent research grant (to E.B.), National Research Initiative Competitive Grants Program (grant 2006-03434 to M.G.I.), the Odysseus program of the Research Foundation-Flanders (J.F.), the Dirección General de Asuntos del Personal Académico – Programa de Apoyo a Proyectos de Investigación e Innovación Tecnológica, Universidad Nacional Autónoma de México (grant IN212509 to S.S. and grants IN225906 and IN212009 to J.G.D.), Consejo Nacional de Ciencia y Technología, Mexico (grant 79736 to S.S. and grant 49267 to J.G.D.), and Programa de Apoyos para la Superación del Personal Académico – Universidad Nacional Autónoma de México (J.G.D.).

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  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1 Auxin-response gradient in the central cylinder of the distal root portion of the DR5rev::GFP Arabidopsis line.

Fig. S2 Restoration of IAA transport after 1-naphthylphthalamic acid (NPA) treatment.

Fig. S3 Experimental setup to evidence the irreversible nature of transient inhibition of lateral root initiation in Arabidopsis using pIAA14::mIAA14-GR line.

Fig. S4 Gradient in probability of pericycle cells to initiate lateral roots upon local auxin treatment in Arabidopsis CYCB1;1DB::GUS line.

Fig. S5 Dynamics of pericycle cell activation after auxin treatment.

Fig. S6 Gradient in probability of pericycle cells to initiate lateral roots in tomato plants.

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