Formation of a subero-lignified apical deposit in root tip of radish (Raphanus sativus) as a response to copper stress

Formation of a subero-lignified apical deposit in root tip of radish (Raphanus sativus) as a... Abstract Background and Aims Heavy metals induce changes in root metabolism and physiology, which can lead to a complex remodelling of the root system. The final morphological responses of radish (Raphanus sativus) roots exposed to toxic concentrations of the heavy metal (Cu) include root growth inhibition, differentiation of xylem vessels close to the root tip, enhanced suberin lamellae deposition and enhanced lateral root production. Recently, we have found that such changes in root morphology and anatomy are coupled to the formation of a subero-lignified apical deposit (SLAD) very close to the root tip. Methods To clarify the details of the formation of a SLAD in the root tip, we conducted experiments with radish roots exposed to a high Cu concentration (60 µm). Histochemical analysis of lignin and suberin as well as analysis of spatial–temporal characteristics of SLAD formation were performed by bright-field, fluorescence and confocal microscopy. Key Results This unique structure, not longer than 100 µm, consists of modified cell walls of the central cylinder that are encircled by a short cylinder of prematurely suberized endodermal cells. A SLAD starts to form, in both primary and lateral roots, after cessation of root elongation, and it is coupled with xylem differentiation and root branching close to the root apex. We noticed that deposition of phenolic substances into a SLAD, mainly suberin in the endodermis, is spatially separated from suberization or lignification in basally located endodermis. Conclusions Although the main reason for formation of a SLAD is elusive, we suggest that it is a part of stress-induced responses which relate to decreased root growth or permeability in heavy metal stress. Apex, central cylinder, copper, endodermis, lignin, subero-lignified apical deposit (SLAD), radish (Raphanus sativus L.), root anatomy, root transition zone, suberin INTRODUCTION Deposition of various kinds of phenolic compounds that are structural components of suberin and lignin in cell walls is a common mechanism for protection of plant tissues and organs. These metabolites prevent damage caused by gravity, wind or pathogens. In the below-ground tissues, more stressed by conditions and the composition of substrate (soil), they play an important role in management of water and nutrient deficiencies or redundancies. In the central part of the root, i.e. the endodermis and central cylinder, production of phenolics and their deposition into cell walls are a very important part of uptake and transport management and regulation (Bernards, 2002; Enstone et al., 2003; Voxeur et al., 2015). Suberin, although not solely a phenolic substance, is chemically a hydrophobic polyester biopolymer composed of aliphatic and aromatic domains, that in turn are composed of phenolic domains (Ranathunge et al., 2011). The alterations of these domains give rise to characteristic dark and bright layers, typically seen by electron microscopic visualization of suberin. While aliphatic polyesters of glycerol with fatty acids and their derivatives cause emergence of light bands, aromatics, mainly ferulic acid and other phenolics, cause the appearance of dark bands (Graça and Santos, 2007; Vishwanath et al., 2015). The polymeric character of suberin allows three-dimensional organization of its constituents and the resulting matrix non-covalently binds waxes, causing hydrophobicity and impermeability of suberized cell walls to water (Bernards, 2002; Schreiber, 2010; Vishwanath et al., 2015), gasses (Soukup et al., 2007) or pathogens (Thomas et al., 2007). Conversely, suberized cell walls also decrease leakage and loss of accumulated solutes or gasses out of the root system (Soukup et al., 2007; Watanabe et al., 2013). Suberin is mainly present at the interface between two tissues (Vishwanath et al., 2015) or at sites where a plant builds a barrier facing external conditions (Kolattukudy, 2001; Franke and Schreiber, 2007). In the case of suberin deposition, it is obvious that the process is regulated developmentally (Andersen et al., 2015) but, due to its barrier role, the occurrence or amount of suberin may be strongly affected by environmental conditions. There are many examples where cells start to produce phenolic compounds and suberize earlier due to unfavourable external conditions. For example, in cotton (Gossypium hirsutum L.) seedling roots, high salinity induces suberization of cortical cells at root–shoot connections, which is not present in non-treated roots (Reinhardt and Rost, 1995). A suberized peridermal layer in roots of a monocot, Mervilla plumbea (Lindl.) Speta, also occurs only as a reaction to cadmium (Lux et al. 2010). In waterlogged plants, deposition of suberin in the exodermis is almost an independent feature regardless of oxygen content, contrary to non-wetland species developing suberization in relation to water and the oxygen level (Soukup et al., 2007). Vaculík et al. (2012) described different exo- and endodermal suberin deposition in two willow (Salix caprea L.) isolates based on their origin from contaminated and uncontaminated sites, and more advanced suberization related to cadmium tolerance was also seen in several willow cultivars (Lux et al., 2004). The colonization of bean roots by bacteria leads to formation of a nodule possessing two types of endodermis, both with suberin lamellae. The first type of endodermis envelops a whole nodule and the second type is around vascular tissues (Lotocka, 2007). Apart from well-known exo- and endodermis suberin deposition, there are also known specific deposits of these polyphenols into the epidermis in the form of a non-lamellar diffuse suberin. Additionally, there is a good evidence of environmentally induced deposition of suberin in cells occurring in healing wound tissue (Moon et al., 1984; Thomas et al., 2007; Meyer and Peterson, 2011). Lignin, unlike suberin, is mainly involved in cell and tissue strengthening. Its polymer structure is composed of various phenols, mainly p-coumaryl, coniferyl and sinapyl alcohols. Polymerization of these phenols through oxidative radicalization and following radical coupling gives rise to a heterogeneous lignin structure (Vanholme et al., 2010). Lignin deposition stiffens the cell wall and make it impermeable, but it is often associated with cell death (Voxeur et al., 2015). During development of tissues, lignification is regarded as a final step in cell differentiation (Zhou et al., 2015) and, like suberization, developmentally determined lignification can be enhanced by stress-induced lignification of disturbed tissues (Caño-Delgado et al., 2003; Voxeur et al., 2015). A shift in the normal development of xylem lignification to early deposition of phenolics into metaxylem vessels occurs in the elongation zone of maize roots subjected to water deficit. Such a change is connected to increased transcription of two genes involved in lignin synthesis and positive lignin staining in the root elongation zone. Spatially localized changes in deposition of phenolics is involved in inhibition of wall extensibility and root growth (Fan et al., 2006). The elongation zone of cortical cells is also affected by increased lignification in aluminium (Al)-treated wheat roots. The reaction in plants is variable and dependent on the wheat cultivar and Al concentration (Sasaki et al., 1996). Increased lignification of endodermal cells and xylem vessels is also a reaction of Arabidopsis thaliana (L.) Heynh. roots to copper (Cu) stress (Lequeux et al., 2010). In our previous experiments, we investigated the effect of Cu excess on anatomical characteristics of radish roots with special emphasis on formation of lateral roots and the deposition of lignin and suberin within the roots. During our investigation, we found the presence of a specific deposit inside the root apex that showed enrichment in suberin and lignin. With a lack of knowledge of this phenomenon, we named it a ‘subero-lignified apical deposit’ or ‘SLAD’. Therefore, the aims of the present study are focused on the clarification of details of the SLAD phenomenon. MATERIALS AND METHODS Plant cultivation Radish seeds (Raphanus sativus Clemens F1) were pre-cultivated on moist filter paper in the dark at 25 °C for 4 d. Seedlings were transferred to aerated half-strength Hoagland solution (Hoagland and Arnon, 1950) and cultivated for 3 d until lateral roots formed. Afterwards, the solution was changed and seedlings were divided into two groups: (1) plants cultivated in half-strength Hoagland solution (control) with 0.16 µm CuSO4; and (2) plants treated with half-strength Hoagland solution enriched by 60 µm CuSO4 (Cu 60). Plants were cultivated with a 16 h photoperiod at 25 °C in dark plastic 3 L pots for an additional 96 h. Evaluation of plant production parameters Radish plants growing in the control and Cu treatment were harvested every day (n = 6); root and shoot fresh weight and the length of the primary root were measured in both groups and statistically analysed. Histochemical analysis of root tissues To investigate root anatomy, roots (n = 10) were fixed in 99 % methanol (24 h/4 °C). Afterwards the roots were rinsed with distilled water, cleared for 3 h in a beaker with 10 mL of 80 % lactic acid heated to 70 °C, and observed. For histochemical analysis of suberin and lignin, phloroglucinol [2 % (w/v) in 25 % HCl] and Sudan Red 7B [0.1 % (Sigma-Aldrich) in polyethylene glycol (PEG) (Brundrett et al. 1991); SR 7B] staining were carried out after previous clearing by lactic acid. For staining with Fluorol Yellow 088 [0.01% (w/v) (Sigma-Aldrich); FY088], the dye was dissolved in 80 % lactic acid and roots were subsequently cleared and stained in one step according to Lux et al. (2015). All stained roots were washed in distilled water and mounted in 0.1 % (w/v) FeCl3 solution in 50 % (v/v) glycerol. To calculate the number of SLADs in individual control or Cu 60-treated primary and lateral roots, all roots were fixed and stained with FY088 in lactic acid. The number of roots with or without a SLAD was calculated. During calculation, we considered as a lateral root any secondary root which emerged above the primary root surface. For measurement of SLAD position alongside the root axis, roots (n = 6) were fixed and cleared in lactic acid. Subsequently the cell length of endodermal cells was measured from the endodermal initial cell up to the fully elongated endodermal cell in each root. Root samples for cross-sectioning (n = 4) were fixed in tissue freezing medium (Leica Biosystems) and frozen to –20 °C. Subsequently, the samples were cut into 20 µm thick cross-sections. Sections designated for lignin staining were immersed in phloroglucinol/HCl solution and stained for 1 min. Sections designated for suberin staining were immersed in 0.1 % FY088 in 80 % lactic acid and stained in the dark for 1 h. All stained sections were washed in distilled water and observed. For 3-D reconstruction of SLAD-affected cells, Cu-treated roots were fixed and stained with FY088 dissolved in lactic acid and observed using a confocal microscope. All roots and cross-sections were visualized using a Zeiss Axioscope 2 plus (Zeiss, Jena, Germany) or a Leica M165FC stereomicroscope, both equipped with a UV lamp (FY088, Zeiss set 25 or DsRed, respectively). Pictures were taken with an Olympus DP-72 digital camera. For confocal microscopy, a Leica TCS SP5 (Leica, Wetzlar, Germany) DM-6000 CS microscope with a UV diode, a ×20 or ×60 glycerol immersion lens and Z-stack function was used. The excitation and emission wavelengths were set to 405 and 450–550 nm, respectively. Cell length measurement and Z-stack processing were done in ImageJ software v.1.50i (https://imagej.nih.gov/ij/). Evaluation of the effect of locally applied Cu on SLAD formation To find out whether local application of Cu to the apical part of the root is sufficient to induce SLAD formation or whether the whole root should be exposed to Cu, we conducted new experiments. Young radish seedlings (4 d old), germinated on a wet filter paper, were placed between two wet filter papers, except for the leaves which protruded into the air and the youngest (0.5–1 cm) root tip parts which were placed between another two different wet filter papers. Papers covering the root tip and the rest of the root were separated by a small gap. In the case of control roots, solution without Cu was applied to both parts, i.e. the root tip and the rest of the root. In the case of Cu treatment, the root tip was continuously exposed to Cu 60 treatment and control solution was applied to the rest of the root. Seedlings (n = 6) were fixed every 12 h during continuous 48 h cultivation, and anatomical changes were observed after clearing and staining with FY088 in lactic acid. Analysis of spatial–temporal characteristics of SLAD formation in radish roots To better explain the spatial–temporal changes in root anatomy due to excess Cu, in particular the duration of the incubation period needed to induce SLAD formation in roots and the differences in SLAD formation in lateral roots branched on apical or basal root parts, we conducted an additional experiment. Seven-day-old plants pre-cultivated as described before were separated into ten groups (1–10) and exposed to Cu treatment. After various durations of Cu exposure (characteristic for each individual group), plants (n = 4 for each group) were rinsed with distilled water and transferred back to control solution without Cu for the rest of the cultivation period to recover. The detailed exposure and recovery time for each group is shown in Table 1. Finally, 84 h after the start of treatment, all plants were harvested and fixed, except group 10 which continued growing for an additional 84 h. In all plants, the total number of lateral roots with or without a SLAD was evaluated after clearing and staining with FY088 in lactic acid. Table 1. Scheme of exposure of ten groups of radish plants to Cu 60 treatment followed by return to control solution (recovery) Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  View Large Table 1. Scheme of exposure of ten groups of radish plants to Cu 60 treatment followed by return to control solution (recovery) Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  View Large Statistical analysis In all experiments, a sufficient number of biological replicates were analysed. To make this manuscript more user-friendly, the number of individual biological replicates is mentioned when describing the individual experiments above. Statistical significance was assessed with Student’s t-test using the Statgraphics Centurion XV v. 15.2.05 (StatPoint, Inc., Warrenton, VA, USA) and Excel (Microsoft Office 2007) programs, and a single-step multiple comparisons of means was performed via Tukey test. A P-value <0.05 was defined as significant. RESULTS Effect of Cu excess on plant growth Radish plants exposed to 60 µm Cu displayed visible symptoms of metal toxicity. Green biomass production and root length of treated plants differed greatly from those of the control, non-treated plants. Application of Cu stopped any primary root elongation activity (Fig. 1A) and leaf biomass production (Fig. 1B). In addition, the number of lateral roots was increased, and after 48 h it was significantly higher in Cu-exposed plants than in the controls (Fig. 1C). Fig. 1. View largeDownload slide Plant growth parameters. (A) Root length, (B) shoot fresh weight, and (C) lateral root production of control and Cu 60 treated plants. Data are presented as means ± s.e.; an asterisk indicates significant differences among the treatments at P < 0.05. Fig. 1. View largeDownload slide Plant growth parameters. (A) Root length, (B) shoot fresh weight, and (C) lateral root production of control and Cu 60 treated plants. Data are presented as means ± s.e.; an asterisk indicates significant differences among the treatments at P < 0.05. Radish root anatomy after Cu application As observations revealed, in contrast to the control (Fig. 2A, C), lateral roots and xylem vessels in Cu-affected roots appear to be very close to the root tip (Fig. 2B). Elongated vessel elements in control roots differed from very short xylem elements in treated roots (Fig. 2D). Additionally, in Cu-treated roots, a dark, compact and opaque deposit was localized between xylem vessels and the root tip (Fig. 2B, D). This deposit was completely lacking in control roots. Fig. 2. View largeDownload slide Anatomy and histochemistry of (A, C) control and (B, D–N) Cu-affected roots. (A) Bright field image of a living control root without SLAD formation and (B) a living root treated with Cu forming a SLAD visible as a dark patch marked by an arrowhead. Note the proximity of the lateral root (asterisk) and xylem vessel (double arrowhead) to the root tip. (C) Meristematic zone of cleared control root without differentiated xylem vessels or SLAD formation. (D) The same zone as in (C) after treatment with Cu. The SLAD is visible as a dark patch marked by the arrowhead. Note the short xylem vessel elements (double arrow). (E and F) Cu-affected roots stained on lignin with phloroglucinol/HCl shown as (E) a whole root and (F) a cross-section through the area marked in (E) with an arrowhead. (G) Root sample treated with Cu stained on suberin with Sudan Red 7B. The positive signal comes from a group of cells in the root tip marked by the arrowhead. (H and I) Control roots stained on suberin with Fluorol Yellow 088 as (H) a whole root and (I) a cross-section. (J and K) Cu-treated roots stained on suberin with Fluorol Yellow 088 shown as (J) a whole root and (K) a cross-section through area the marked by an arrowhead. (L) Detail of the cross-section from (K). (M) Confocal imaging of the affected apical parts of Cu-treated roots after Fluorol Yellow 088 staining. (N) Three-dimensional reconstruction of phenolic-rich cell walls of the SLAD in Cu- affected roots. Abbreviations: rc, root cap; rh, rhizodermis; cx, cortex; en, endodermis; cc, central cylinder. Scale bars (A, B) = 200 µm; (C–K) = 100 µm; (L, M) = 20 µm. Fig. 2. View largeDownload slide Anatomy and histochemistry of (A, C) control and (B, D–N) Cu-affected roots. (A) Bright field image of a living control root without SLAD formation and (B) a living root treated with Cu forming a SLAD visible as a dark patch marked by an arrowhead. Note the proximity of the lateral root (asterisk) and xylem vessel (double arrowhead) to the root tip. (C) Meristematic zone of cleared control root without differentiated xylem vessels or SLAD formation. (D) The same zone as in (C) after treatment with Cu. The SLAD is visible as a dark patch marked by the arrowhead. Note the short xylem vessel elements (double arrow). (E and F) Cu-affected roots stained on lignin with phloroglucinol/HCl shown as (E) a whole root and (F) a cross-section through the area marked in (E) with an arrowhead. (G) Root sample treated with Cu stained on suberin with Sudan Red 7B. The positive signal comes from a group of cells in the root tip marked by the arrowhead. (H and I) Control roots stained on suberin with Fluorol Yellow 088 as (H) a whole root and (I) a cross-section. (J and K) Cu-treated roots stained on suberin with Fluorol Yellow 088 shown as (J) a whole root and (K) a cross-section through area the marked by an arrowhead. (L) Detail of the cross-section from (K). (M) Confocal imaging of the affected apical parts of Cu-treated roots after Fluorol Yellow 088 staining. (N) Three-dimensional reconstruction of phenolic-rich cell walls of the SLAD in Cu- affected roots. Abbreviations: rc, root cap; rh, rhizodermis; cx, cortex; en, endodermis; cc, central cylinder. Scale bars (A, B) = 200 µm; (C–K) = 100 µm; (L, M) = 20 µm. Histochemical analysis and characterization of the root tip deposit Cell walls in stressed tissues are characterized by accumulation of lignin and suberin. We identified the presence of these substances in Cu-treated roots, both in whole roots and in cross-sectionss. Phloroglucinol/HCl staining of the whole root revealed the presence of lignin in a small area in the root tip. On cross-sections, the same staining was localized inside the endodermis and central cylinder (Fig. 2E, F). Staining for suberin with SR 7B also revealed pinkish-red staining in the same part of the root, between xylem vessels and the root tip (Fig. 2G). Similarly, FY088 staining for suberin (Fig. 2H–L) was responsible for strong fluorescence in the same area, visible both in the whole root (Fig. 2J) and in cross-sections (Fig. 2K). On cross-sections, similarly to lignin staining, the highest fluorescent signal originated from the endodermis and central cylinder (Fig. 2L). In control roots, such positive staining was completely absent (Fig. 2H, I). Deposition of phenolics, mainly suberin and lignin, in the endodermis and central cylinder was also revealed by confocal imaging and 3-D reconstruction of the affected area. Phenolics were mainly in cell walls in a short segment of endodermal cells and cells of the central cylinder. These subero-lignified structures in both tissues were localized in an area a few micrometres long, positioned approx. 150–250 µm from the root tip. Affected cells of the endodermis were short, in contrast to elongated cells of the central cylinder. All cells in the central cylinder were characterized by deposition of lignin; none of them showed thickenings typical for xylem vessels or non-uniform deposition of phenolics in cell walls (Fig. 2M, N). Due to the very specific localization in the root apex as well as the dominant occurrence of suberin and lignin in cell walls of this structure we propose to name it a ‘subero-lignified apical deposit’ (SLAD). Measurement of the SLAD position based on the length of endodermal cells along the root axis showed that the SLAD is always present in an area with short, non-elongated endodermal cells, which is characterized by transition between the meristematic and elongation zone. In Cu-treated roots with a formed SLAD, the position of the transition zone is similar to that in control roots, and the length of elongated cells in the elongation zone is lower (Fig. 3). Fig. 3. View largeDownload slide Relationship between cell order from the root initial cells (quiescence centre) and cell length in control (dashed line) and Cu-treated roots (solid line). The grey zone defines cells in treated roots where a SLAD can be formed. Fig. 3. View largeDownload slide Relationship between cell order from the root initial cells (quiescence centre) and cell length in control (dashed line) and Cu-treated roots (solid line). The grey zone defines cells in treated roots where a SLAD can be formed. Evaluation of the effect of locally applied Cu on SLAD formation Roots treated with Cu solely in the root tip region were harvested every 12 h for 2 d and anatomical features were observed. In the control treatment, the root apex did not show any changes in root anatomy, shift of xylem vessels or lateral roots closer to the root apex, or SLAD formation even after 48 h of treatment. However, roots treated with Cu showed symptoms of toxicity. After 12 h of Cu treatment, xylem vessels started to differentiate closer to the root apex but lateral roots or a SLAD were not visible in this zone. After 24 h of Cu treatment, xylem vessels and lateral roots continuously shifted their differentiation and appeared closer to the root apex than in control roots, although a SLAD was still not formed. After 36 h, xylem vessels ceased differentiation and were localized at a very small distance from the root tip. Also, lateral roots appeared closer to the root apex and were visible as small bulges. At this stage, a SLAD was formed as a group of cells stained by FY088. After 48 h, SLAD and xylem vessel differentiation were not different from the previous stage, which indicated cessation of primary root growth, contrary to continuously emerging and growing lateral root primordia (Fig. 4). Fig. 4. View largeDownload slide Schematic representation of the root anatomy influenced by a local application of Cu to a small part of the root tip. In the time course, xylem vessels and lateral roots differentiate closer to the root apex. After 36 h of Cu exposure, SLAD formation occurs. After 48 h, xylem vessels and the SLAD occupy the root tip and stop differentiation, in contrast to continuously growing lateral roots. Fig. 4. View largeDownload slide Schematic representation of the root anatomy influenced by a local application of Cu to a small part of the root tip. In the time course, xylem vessels and lateral roots differentiate closer to the root apex. After 36 h of Cu exposure, SLAD formation occurs. After 48 h, xylem vessels and the SLAD occupy the root tip and stop differentiation, in contrast to continuously growing lateral roots. Analysis of spatial–temporal characteristics of SLAD formation in radish roots The relationship between SLAD formation and Cu exposure time was investigated in a recovery experiment. Ten groups of plants (1–10) were exposed to various durations of Cu 60 treatment and then subsequently transferred to control media for recovery (Table 1). Root morphology changes and SLAD distribution in both primary and lateral roots were calculated. In group 1, i.e. plants growing for the whole experimental period (84 h) in a control solution, almost all lateral roots branched at the basal part of the primary root and the total number of laterals decreased towards the apical part. Both primary and lateral control roots were completely without any SLAD in their apices (Fig. 5A). Plants in groups 2 and 3, exposed to Cu for 1 and 12 h, respectively, had shifted lateral root formation towards the primary root apex. However, the apex of the primary root was completely lacking any SLAD. On the other hand, a SLAD was formed in 9 % of lateral roots after 1 h exposure and in 20 % after 12 h exposure. In both groups, all SLADs forming laterals were localized at the basal part of the primary root (Fig. 5B). Moreover, some lateral roots showed a region close to the apex, where xylem vessels were short and stacked but without a formed SLAD (data not shown). Because this region was followed by elongated xylem vessels, it possibly indicates a partial decrease in root growth during Cu exposure followed by new growth without SLAD formation. Plants in groups 4–9, exposed to Cu from 24 to 84 h, showed typical morphological changes induced by metal toxicity. In contrast to groups 2 and 3, the primary roots in groups 4–9 always formed a SLAD in their apices and the distribution of lateral roots was more shifted towards their tips. In groups 4–9, a total of 34–53 % of lateral roots formed a SLAD in their apices. The formation of a SLAD was the most prominent in those lateral roots that were developed close to the primary root base and decreased towards the primary root apex. The laterals branched at the apical part of the primary root did not form any SLAD (Fig. 5C). Group 10, fully exposed to Cu for 168 h without any recovery, showed a SLAD in the primary root apex, and nearly 70 % of all lateral roots also formed a SLAD (Fig. 5D). Moreover, deposition of suberin into SLADs in lateral roots also affected other endodermal cells and was visible as a differentiated endodermis along the whole lateral root axis (Fig. 5E). Fig. 5. View largeDownload slide Root morphological changes and distribution of SLAD-affected roots (red root) and roots without a SLAD (black root) after the recovery experiment. Percentages show the number of SLAD-affected lateral roots in individual groups. Root lengths on the schemes are illustrative, but the distribution and proportion of lateral roots with or without a SLAD depict the real situation. (A) Scheme of a control root (group 1), fully exposed to control solution, that does not form a SLAD in laterals or the main root. (B) Scheme of morphological changes of a root after 12 h of Cu exposure followed by a 72 h recovery time (group 3). Note the lack of SLAD formation in the primary root, shifted lateral root production towards the root apex and formation of a SLAD in 20 % of lateral roots. (C) Scheme of morphological changes of a root after 72 h of Cu exposure followed by a 12 h recovery time (group 8). A SLAD is formed in the primary root and in 45 % of all lateral roots. (D) Scheme of root morphology after a 168 h exposure to Cu (group 10). Around 70 % of all lateral roots form a SLAD, and a SLAD is also present in the primary root. (E) A lateral root (LR) after a 168 h exposure to Cu (group 10) and FY088 staining. The presence of suberin is visible in the SLAD (arrowhead) and in all endodermal cells (asterisk) from the SLAD to the junction on the primary root (PR). Fig. 5. View largeDownload slide Root morphological changes and distribution of SLAD-affected roots (red root) and roots without a SLAD (black root) after the recovery experiment. Percentages show the number of SLAD-affected lateral roots in individual groups. Root lengths on the schemes are illustrative, but the distribution and proportion of lateral roots with or without a SLAD depict the real situation. (A) Scheme of a control root (group 1), fully exposed to control solution, that does not form a SLAD in laterals or the main root. (B) Scheme of morphological changes of a root after 12 h of Cu exposure followed by a 72 h recovery time (group 3). Note the lack of SLAD formation in the primary root, shifted lateral root production towards the root apex and formation of a SLAD in 20 % of lateral roots. (C) Scheme of morphological changes of a root after 72 h of Cu exposure followed by a 12 h recovery time (group 8). A SLAD is formed in the primary root and in 45 % of all lateral roots. (D) Scheme of root morphology after a 168 h exposure to Cu (group 10). Around 70 % of all lateral roots form a SLAD, and a SLAD is also present in the primary root. (E) A lateral root (LR) after a 168 h exposure to Cu (group 10) and FY088 staining. The presence of suberin is visible in the SLAD (arrowhead) and in all endodermal cells (asterisk) from the SLAD to the junction on the primary root (PR). DISCUSSION There is knowledge about development of root hairs or lateral roots very close to the apex in the case of a stress-induced morphological response (Potters et al., 2007). Another stress-induced plant response is production and deposition of phenolic substances, e.g. suberin or lignin, into damaged cells as well as enhancement of development of root apoplasmic barriers when plants suffer under hostile conditions (Caño-Delgado et al., 2003; Enstone et al., 2003). Similar symptoms were observed in radish plants exposed to a high concentration of Cu in our experiments. Moreover, we found specific deposition of the phenolics suberin and lignin inside a root tip area that we described as a subero-lignified apical deposit (SLAD). This is in contrast to the well accepted model of root endodermis suberization which is based on continuous differentiation of cells from the root base to the apex. In the case of endodermal suberization, it is not only an environmentally, but also a positionally regulated process. Unilateral exposure of maize roots to a high concentration of Cd caused decreased cell elongation activity in the Cd-exposed part of the root and, simultaneously, cells of this part deposited suberin in cell walls more intensively and closer to the root tip in contrast to the Cd-free root site (Lux et al., 2011). Endodermal cells positioned near the phloem poles start to suberize much closer to the root tip than other endodermal cells, and the number of suberized cells increases toward the base. Suberization of the endodermis is not completed until all endodermal cells in the basal part of the root are completely covered by suberin. The exception is a group of passage cells, located near to the xylem poles that suberize only occasionally, as was shown in garlic (Allium cepa L.) (Waduwara et al., 2008) and soybean [Glycine max (L.) Merr.] roots (Thomas et al., 2007). Previous observations on A. thaliana showed that emerging lateral roots induced suberization in a well-defined group of endodermal cells located above the lateral root primordia or around emerged lateral roots. Such cells may suberize prematurely, very early and during the first stages of lateral root growth, and may protect primary root from an uncontrolled influx of solutes (Martinka et al., 2012). In SLADs, deposition of phenolics into endodermal cells can be separated from the rest of the endodermis. As revealed by the experiment with Cu stress applied solely to the root tip, SLADs can be formed independently and endodermal cells can suberize prematurely in this part of the root, compared with other endodermal cells. Premature suberization of endodermal cells in the endodermal part of SLADs is peculiar for several reasons. According to cell length measurement, a SLAD affects cells in the basal part of the meristematic zone. Normally, young cells in apical meristems are undifferentiated and, for example, in A. thaliana roots, local ablation of an atrichoblast cell in the rhizodermis induces changes in a trichoblast cell located nearby which does not develop into a root hair but takes the position and function of an ablated atrichoblast cell (Berger et al., 1998). The fate of cells in the meristematic zone is not strongly pre-determined. If all meristematic cells are exposed to the same conditions and environment, but only endodermal and central cylinder cells start to produce secondary metabolites such as phenolics, they are probably pre-determined to have such a function very early. In the case of the endodermis, such a reaction is supported by the important position of this layer on the border between internal vascular tissues and the external environment (Robbins et al., 2014). Current knowledge about the very high plasticity of endodermis development and suberin deposition influenced by many signals and environmental conditions can also support these findings (Barberon, 2017). Even older data showed the presence of phenolic-storing cells in plant tissues which react very rapidly to stress and subsequently may become lignified or suberized (e.g. Beckman, 2000, and references therein). One layer characteristic of such cells is the endodermis (Mueller and Beckman, 1976). Pro-endodermis, a layer of young endodermal cells before their differentiation, is in some species also characterized by a higher phenolic content (Beckman, 2000; Ma and Peterson, 2003). Auxin hormonal regulation can also explain early endodermis differentiation during SLAD formation. Auxin transported from the root base to the apex changes direction at the root apex, and local gradients of this hormone induce maintenance of meristematic activity and various reactions and developmental responses (Moubayidin et al., 2009; Overvoorde et al., 2010). If unfavourable conditions change the hormonal concentrations [through degradation of reactive oxygen species or peroxidases by changes of pH, failure in transport or incorrect localization of the cytoskeleton (Pasternak et al., 2005)], the sensitive root tip may react very rapidly and reactions start preferentially in these areas. Deposition of phenolics into the narrow area of the central cylinder encircled by differentiated endodermal cells can be related to differentiation of xylem, even though vessels are not visible in this part of the SLAD. Normally xylem differentiation induces deposition of lignin and cell death in elongated cells in the central cylinder (Srivastava and Singh, 1972; Fukuda, 1997; van Doorn and Woltering, 2005). If initial cells in meristems of A. thaliana roots are under DNA-damaging stress (γ-rays, UV light, heavy metals, reactive oxygen species or toxic compounds), they are selectively eliminated and killed. Because initials produce lineages of cells and genetic information, damage induced by stress can lead to mutation in daughter cells, too (Fulcher and Sablowski, 2009). Therefore, DNA-damaging factors in initials leads to programmed cell death, and dead cells are in the meristematic apical zone observable after propidium iodide staining as intensively stained structures (Furukawa et al., 2010). Also, despite common high susceptibility of all initials to DNA damage, cells of the central cylinder are much more sensitive to damage and cell death (Fulcher and Sablowski, 2009). Therefore, higher susceptibility and possible cell death of cells in the central cylinder might relate to the observed stress-induced deposition of phenolics in our experiments. Radish root morphology affected by Cu, including an increased number of lateral roots, their shift to the root apex and modified deposition of phenolics, is in agreement with Cu-induced reorganization of A. thaliana root architecture (Lequeux et al., 2010). The described morphological changes in roots are similar to stress reactions called stress-induced morphogenic responses (SIMRs) (Potters et al., 2007). According to these authors, in the case of an SIMR, stress does not kill the plant, but causes (1) inhibition of elongation; (2) local stimulation of cell division; and (3) alterations to cell differentiation. Subsequently, a new developmental programme completely changes the morphology of the stressed plant (Potters et al., 2009). Due to this, we assume that the formation of SLADs can be one part of the SIMR reaction, because (1) SLAD-affected roots ceased their elongation activity; (2) SLAD formation occurs in roots along with apically stimulated lateral root growth; and (3) differentiation of cells in SLADs is altered by deposition of suberin and lignin. According to the recovery experiment, 1 h of Cu 60 exposure is enough to induce SLAD formation in some lateral roots, and exposure for >24 h induces SLAD formation in half of all lateral roots regardless of the recovery time. However, if SLAD formation is a part of the SIMR, deposition of phenolics into SLADs is very probably not a reason for but a reaction to limited growth of the root. Because SLAD formation follows and does not precede emergence of laterals in the apex, it is also not responsible for reorganization or remodelling of the root system. It is similar to the reaction of wheat roots to Al stress, when lignification affects sensitive roots. Deposition of phenolics into elongating cells is not a reason for cessation of growth, but a reaction to damage caused by the element in excess (Sasaki et al. 1996). In the case of the endodermis, SLAD-forming deposits are typically located in the cell walls of short cells present just before the onset of elongation. The change from meristematic to elongation activity occurs normally in the transition zone. The transition zone is composed of developmentally plastic cells (Verbelen et al., 2006), and plasticity allows the root to undergoe diverse tropisms and react to internal or external stimuli (Baluška et al., 2010). Changes in root growth in the transition zone are affected by reaction to nitrate (Trevisan et al., 2015), osmotic stress (Baluška and Mancuso, 2013) or Al stress (Sivaguru and Horst, 1998), among others. Cells of the transition zone are rich in photosynthates transported by phloem differentiating just in this portion of the root (Verbelen et al., 2006). The transition zone is also responsible for root growth regulation via phytohormones (Dello Ioio et al., 2007; Ubeda-Tomás et al., 2009). High sensitivity, unloading of metabolites and high oxygen influx despite low meristematic activity in the transition zone (Baluška and Mancuso, 2013) provide suitable conditions for deposition of phenolics and SLAD formation in this zone. The role of the SLAD and deposition of phenolic-containing substances into a narrow area of the transition zone between xylem vessels and root initials in the SIMR reaction may be in exclusion of supplies of sugar and metabolites to the quiescence centre or initial cells to restrict growth in the case of unfavourable conditions. Similarly, the SLAD may prevent xylem vessels from taking up unfavourable solutes. An increased number of lateral roots and their shift to the root apex in the SIMR reaction can compensate for loss of primary root activity but, if unfavourable conditions continue, all lateral roots stop growth and form a SLAD. Moreover, in such negative conditions, phenolics (suberin) are present not only in SLADs, but in all endodermal cells from the root base to the SLAD. Such an isolation of the root from the negative environment by phenolics deposited into SLADs and the endodermis may also explain the formation and existence of SLADs in the root apex. Conclusion Our recent findings revealed the presence of a specific structure formed in the root apex of radish after Cu exposure. This structure is localized in the transition zone of the root and consists of phenolic-enriched components lignin and suberin. Therefore, we named this structure ‘subero-lignified apical deposit’ or SLAD. Formation of a SLAD depends on the time of exposure to high Cu concentrations and affects endodermal and central cylinder cells in both primary and lateral roots. Moreover, it is connected to stress-induced remodelling of the root system, such as a decrease in root growth or an increase in lateral root production. The exact role of a SLAD can only be predicted at present; however, we assume that its presence in the root tip may enhance endodermal barrier function, decrease xylem conductivity or limit phloem transport. 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Formation of a subero-lignified apical deposit in root tip of radish (Raphanus sativus) as a response to copper stress

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Abstract

Abstract Background and Aims Heavy metals induce changes in root metabolism and physiology, which can lead to a complex remodelling of the root system. The final morphological responses of radish (Raphanus sativus) roots exposed to toxic concentrations of the heavy metal (Cu) include root growth inhibition, differentiation of xylem vessels close to the root tip, enhanced suberin lamellae deposition and enhanced lateral root production. Recently, we have found that such changes in root morphology and anatomy are coupled to the formation of a subero-lignified apical deposit (SLAD) very close to the root tip. Methods To clarify the details of the formation of a SLAD in the root tip, we conducted experiments with radish roots exposed to a high Cu concentration (60 µm). Histochemical analysis of lignin and suberin as well as analysis of spatial–temporal characteristics of SLAD formation were performed by bright-field, fluorescence and confocal microscopy. Key Results This unique structure, not longer than 100 µm, consists of modified cell walls of the central cylinder that are encircled by a short cylinder of prematurely suberized endodermal cells. A SLAD starts to form, in both primary and lateral roots, after cessation of root elongation, and it is coupled with xylem differentiation and root branching close to the root apex. We noticed that deposition of phenolic substances into a SLAD, mainly suberin in the endodermis, is spatially separated from suberization or lignification in basally located endodermis. Conclusions Although the main reason for formation of a SLAD is elusive, we suggest that it is a part of stress-induced responses which relate to decreased root growth or permeability in heavy metal stress. Apex, central cylinder, copper, endodermis, lignin, subero-lignified apical deposit (SLAD), radish (Raphanus sativus L.), root anatomy, root transition zone, suberin INTRODUCTION Deposition of various kinds of phenolic compounds that are structural components of suberin and lignin in cell walls is a common mechanism for protection of plant tissues and organs. These metabolites prevent damage caused by gravity, wind or pathogens. In the below-ground tissues, more stressed by conditions and the composition of substrate (soil), they play an important role in management of water and nutrient deficiencies or redundancies. In the central part of the root, i.e. the endodermis and central cylinder, production of phenolics and their deposition into cell walls are a very important part of uptake and transport management and regulation (Bernards, 2002; Enstone et al., 2003; Voxeur et al., 2015). Suberin, although not solely a phenolic substance, is chemically a hydrophobic polyester biopolymer composed of aliphatic and aromatic domains, that in turn are composed of phenolic domains (Ranathunge et al., 2011). The alterations of these domains give rise to characteristic dark and bright layers, typically seen by electron microscopic visualization of suberin. While aliphatic polyesters of glycerol with fatty acids and their derivatives cause emergence of light bands, aromatics, mainly ferulic acid and other phenolics, cause the appearance of dark bands (Graça and Santos, 2007; Vishwanath et al., 2015). The polymeric character of suberin allows three-dimensional organization of its constituents and the resulting matrix non-covalently binds waxes, causing hydrophobicity and impermeability of suberized cell walls to water (Bernards, 2002; Schreiber, 2010; Vishwanath et al., 2015), gasses (Soukup et al., 2007) or pathogens (Thomas et al., 2007). Conversely, suberized cell walls also decrease leakage and loss of accumulated solutes or gasses out of the root system (Soukup et al., 2007; Watanabe et al., 2013). Suberin is mainly present at the interface between two tissues (Vishwanath et al., 2015) or at sites where a plant builds a barrier facing external conditions (Kolattukudy, 2001; Franke and Schreiber, 2007). In the case of suberin deposition, it is obvious that the process is regulated developmentally (Andersen et al., 2015) but, due to its barrier role, the occurrence or amount of suberin may be strongly affected by environmental conditions. There are many examples where cells start to produce phenolic compounds and suberize earlier due to unfavourable external conditions. For example, in cotton (Gossypium hirsutum L.) seedling roots, high salinity induces suberization of cortical cells at root–shoot connections, which is not present in non-treated roots (Reinhardt and Rost, 1995). A suberized peridermal layer in roots of a monocot, Mervilla plumbea (Lindl.) Speta, also occurs only as a reaction to cadmium (Lux et al. 2010). In waterlogged plants, deposition of suberin in the exodermis is almost an independent feature regardless of oxygen content, contrary to non-wetland species developing suberization in relation to water and the oxygen level (Soukup et al., 2007). Vaculík et al. (2012) described different exo- and endodermal suberin deposition in two willow (Salix caprea L.) isolates based on their origin from contaminated and uncontaminated sites, and more advanced suberization related to cadmium tolerance was also seen in several willow cultivars (Lux et al., 2004). The colonization of bean roots by bacteria leads to formation of a nodule possessing two types of endodermis, both with suberin lamellae. The first type of endodermis envelops a whole nodule and the second type is around vascular tissues (Lotocka, 2007). Apart from well-known exo- and endodermis suberin deposition, there are also known specific deposits of these polyphenols into the epidermis in the form of a non-lamellar diffuse suberin. Additionally, there is a good evidence of environmentally induced deposition of suberin in cells occurring in healing wound tissue (Moon et al., 1984; Thomas et al., 2007; Meyer and Peterson, 2011). Lignin, unlike suberin, is mainly involved in cell and tissue strengthening. Its polymer structure is composed of various phenols, mainly p-coumaryl, coniferyl and sinapyl alcohols. Polymerization of these phenols through oxidative radicalization and following radical coupling gives rise to a heterogeneous lignin structure (Vanholme et al., 2010). Lignin deposition stiffens the cell wall and make it impermeable, but it is often associated with cell death (Voxeur et al., 2015). During development of tissues, lignification is regarded as a final step in cell differentiation (Zhou et al., 2015) and, like suberization, developmentally determined lignification can be enhanced by stress-induced lignification of disturbed tissues (Caño-Delgado et al., 2003; Voxeur et al., 2015). A shift in the normal development of xylem lignification to early deposition of phenolics into metaxylem vessels occurs in the elongation zone of maize roots subjected to water deficit. Such a change is connected to increased transcription of two genes involved in lignin synthesis and positive lignin staining in the root elongation zone. Spatially localized changes in deposition of phenolics is involved in inhibition of wall extensibility and root growth (Fan et al., 2006). The elongation zone of cortical cells is also affected by increased lignification in aluminium (Al)-treated wheat roots. The reaction in plants is variable and dependent on the wheat cultivar and Al concentration (Sasaki et al., 1996). Increased lignification of endodermal cells and xylem vessels is also a reaction of Arabidopsis thaliana (L.) Heynh. roots to copper (Cu) stress (Lequeux et al., 2010). In our previous experiments, we investigated the effect of Cu excess on anatomical characteristics of radish roots with special emphasis on formation of lateral roots and the deposition of lignin and suberin within the roots. During our investigation, we found the presence of a specific deposit inside the root apex that showed enrichment in suberin and lignin. With a lack of knowledge of this phenomenon, we named it a ‘subero-lignified apical deposit’ or ‘SLAD’. Therefore, the aims of the present study are focused on the clarification of details of the SLAD phenomenon. MATERIALS AND METHODS Plant cultivation Radish seeds (Raphanus sativus Clemens F1) were pre-cultivated on moist filter paper in the dark at 25 °C for 4 d. Seedlings were transferred to aerated half-strength Hoagland solution (Hoagland and Arnon, 1950) and cultivated for 3 d until lateral roots formed. Afterwards, the solution was changed and seedlings were divided into two groups: (1) plants cultivated in half-strength Hoagland solution (control) with 0.16 µm CuSO4; and (2) plants treated with half-strength Hoagland solution enriched by 60 µm CuSO4 (Cu 60). Plants were cultivated with a 16 h photoperiod at 25 °C in dark plastic 3 L pots for an additional 96 h. Evaluation of plant production parameters Radish plants growing in the control and Cu treatment were harvested every day (n = 6); root and shoot fresh weight and the length of the primary root were measured in both groups and statistically analysed. Histochemical analysis of root tissues To investigate root anatomy, roots (n = 10) were fixed in 99 % methanol (24 h/4 °C). Afterwards the roots were rinsed with distilled water, cleared for 3 h in a beaker with 10 mL of 80 % lactic acid heated to 70 °C, and observed. For histochemical analysis of suberin and lignin, phloroglucinol [2 % (w/v) in 25 % HCl] and Sudan Red 7B [0.1 % (Sigma-Aldrich) in polyethylene glycol (PEG) (Brundrett et al. 1991); SR 7B] staining were carried out after previous clearing by lactic acid. For staining with Fluorol Yellow 088 [0.01% (w/v) (Sigma-Aldrich); FY088], the dye was dissolved in 80 % lactic acid and roots were subsequently cleared and stained in one step according to Lux et al. (2015). All stained roots were washed in distilled water and mounted in 0.1 % (w/v) FeCl3 solution in 50 % (v/v) glycerol. To calculate the number of SLADs in individual control or Cu 60-treated primary and lateral roots, all roots were fixed and stained with FY088 in lactic acid. The number of roots with or without a SLAD was calculated. During calculation, we considered as a lateral root any secondary root which emerged above the primary root surface. For measurement of SLAD position alongside the root axis, roots (n = 6) were fixed and cleared in lactic acid. Subsequently the cell length of endodermal cells was measured from the endodermal initial cell up to the fully elongated endodermal cell in each root. Root samples for cross-sectioning (n = 4) were fixed in tissue freezing medium (Leica Biosystems) and frozen to –20 °C. Subsequently, the samples were cut into 20 µm thick cross-sections. Sections designated for lignin staining were immersed in phloroglucinol/HCl solution and stained for 1 min. Sections designated for suberin staining were immersed in 0.1 % FY088 in 80 % lactic acid and stained in the dark for 1 h. All stained sections were washed in distilled water and observed. For 3-D reconstruction of SLAD-affected cells, Cu-treated roots were fixed and stained with FY088 dissolved in lactic acid and observed using a confocal microscope. All roots and cross-sections were visualized using a Zeiss Axioscope 2 plus (Zeiss, Jena, Germany) or a Leica M165FC stereomicroscope, both equipped with a UV lamp (FY088, Zeiss set 25 or DsRed, respectively). Pictures were taken with an Olympus DP-72 digital camera. For confocal microscopy, a Leica TCS SP5 (Leica, Wetzlar, Germany) DM-6000 CS microscope with a UV diode, a ×20 or ×60 glycerol immersion lens and Z-stack function was used. The excitation and emission wavelengths were set to 405 and 450–550 nm, respectively. Cell length measurement and Z-stack processing were done in ImageJ software v.1.50i (https://imagej.nih.gov/ij/). Evaluation of the effect of locally applied Cu on SLAD formation To find out whether local application of Cu to the apical part of the root is sufficient to induce SLAD formation or whether the whole root should be exposed to Cu, we conducted new experiments. Young radish seedlings (4 d old), germinated on a wet filter paper, were placed between two wet filter papers, except for the leaves which protruded into the air and the youngest (0.5–1 cm) root tip parts which were placed between another two different wet filter papers. Papers covering the root tip and the rest of the root were separated by a small gap. In the case of control roots, solution without Cu was applied to both parts, i.e. the root tip and the rest of the root. In the case of Cu treatment, the root tip was continuously exposed to Cu 60 treatment and control solution was applied to the rest of the root. Seedlings (n = 6) were fixed every 12 h during continuous 48 h cultivation, and anatomical changes were observed after clearing and staining with FY088 in lactic acid. Analysis of spatial–temporal characteristics of SLAD formation in radish roots To better explain the spatial–temporal changes in root anatomy due to excess Cu, in particular the duration of the incubation period needed to induce SLAD formation in roots and the differences in SLAD formation in lateral roots branched on apical or basal root parts, we conducted an additional experiment. Seven-day-old plants pre-cultivated as described before were separated into ten groups (1–10) and exposed to Cu treatment. After various durations of Cu exposure (characteristic for each individual group), plants (n = 4 for each group) were rinsed with distilled water and transferred back to control solution without Cu for the rest of the cultivation period to recover. The detailed exposure and recovery time for each group is shown in Table 1. Finally, 84 h after the start of treatment, all plants were harvested and fixed, except group 10 which continued growing for an additional 84 h. In all plants, the total number of lateral roots with or without a SLAD was evaluated after clearing and staining with FY088 in lactic acid. Table 1. Scheme of exposure of ten groups of radish plants to Cu 60 treatment followed by return to control solution (recovery) Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  View Large Table 1. Scheme of exposure of ten groups of radish plants to Cu 60 treatment followed by return to control solution (recovery) Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  Group  1  2  3  4  5  6  7  8  9  10  Time in Cu 60 solution (h)  0  1  12  24  36  48  60  72  84  168  Time in control solution (h)  84  83  72  60  48  36  24  12  0  0  View Large Statistical analysis In all experiments, a sufficient number of biological replicates were analysed. To make this manuscript more user-friendly, the number of individual biological replicates is mentioned when describing the individual experiments above. Statistical significance was assessed with Student’s t-test using the Statgraphics Centurion XV v. 15.2.05 (StatPoint, Inc., Warrenton, VA, USA) and Excel (Microsoft Office 2007) programs, and a single-step multiple comparisons of means was performed via Tukey test. A P-value <0.05 was defined as significant. RESULTS Effect of Cu excess on plant growth Radish plants exposed to 60 µm Cu displayed visible symptoms of metal toxicity. Green biomass production and root length of treated plants differed greatly from those of the control, non-treated plants. Application of Cu stopped any primary root elongation activity (Fig. 1A) and leaf biomass production (Fig. 1B). In addition, the number of lateral roots was increased, and after 48 h it was significantly higher in Cu-exposed plants than in the controls (Fig. 1C). Fig. 1. View largeDownload slide Plant growth parameters. (A) Root length, (B) shoot fresh weight, and (C) lateral root production of control and Cu 60 treated plants. Data are presented as means ± s.e.; an asterisk indicates significant differences among the treatments at P < 0.05. Fig. 1. View largeDownload slide Plant growth parameters. (A) Root length, (B) shoot fresh weight, and (C) lateral root production of control and Cu 60 treated plants. Data are presented as means ± s.e.; an asterisk indicates significant differences among the treatments at P < 0.05. Radish root anatomy after Cu application As observations revealed, in contrast to the control (Fig. 2A, C), lateral roots and xylem vessels in Cu-affected roots appear to be very close to the root tip (Fig. 2B). Elongated vessel elements in control roots differed from very short xylem elements in treated roots (Fig. 2D). Additionally, in Cu-treated roots, a dark, compact and opaque deposit was localized between xylem vessels and the root tip (Fig. 2B, D). This deposit was completely lacking in control roots. Fig. 2. View largeDownload slide Anatomy and histochemistry of (A, C) control and (B, D–N) Cu-affected roots. (A) Bright field image of a living control root without SLAD formation and (B) a living root treated with Cu forming a SLAD visible as a dark patch marked by an arrowhead. Note the proximity of the lateral root (asterisk) and xylem vessel (double arrowhead) to the root tip. (C) Meristematic zone of cleared control root without differentiated xylem vessels or SLAD formation. (D) The same zone as in (C) after treatment with Cu. The SLAD is visible as a dark patch marked by the arrowhead. Note the short xylem vessel elements (double arrow). (E and F) Cu-affected roots stained on lignin with phloroglucinol/HCl shown as (E) a whole root and (F) a cross-section through the area marked in (E) with an arrowhead. (G) Root sample treated with Cu stained on suberin with Sudan Red 7B. The positive signal comes from a group of cells in the root tip marked by the arrowhead. (H and I) Control roots stained on suberin with Fluorol Yellow 088 as (H) a whole root and (I) a cross-section. (J and K) Cu-treated roots stained on suberin with Fluorol Yellow 088 shown as (J) a whole root and (K) a cross-section through area the marked by an arrowhead. (L) Detail of the cross-section from (K). (M) Confocal imaging of the affected apical parts of Cu-treated roots after Fluorol Yellow 088 staining. (N) Three-dimensional reconstruction of phenolic-rich cell walls of the SLAD in Cu- affected roots. Abbreviations: rc, root cap; rh, rhizodermis; cx, cortex; en, endodermis; cc, central cylinder. Scale bars (A, B) = 200 µm; (C–K) = 100 µm; (L, M) = 20 µm. Fig. 2. View largeDownload slide Anatomy and histochemistry of (A, C) control and (B, D–N) Cu-affected roots. (A) Bright field image of a living control root without SLAD formation and (B) a living root treated with Cu forming a SLAD visible as a dark patch marked by an arrowhead. Note the proximity of the lateral root (asterisk) and xylem vessel (double arrowhead) to the root tip. (C) Meristematic zone of cleared control root without differentiated xylem vessels or SLAD formation. (D) The same zone as in (C) after treatment with Cu. The SLAD is visible as a dark patch marked by the arrowhead. Note the short xylem vessel elements (double arrow). (E and F) Cu-affected roots stained on lignin with phloroglucinol/HCl shown as (E) a whole root and (F) a cross-section through the area marked in (E) with an arrowhead. (G) Root sample treated with Cu stained on suberin with Sudan Red 7B. The positive signal comes from a group of cells in the root tip marked by the arrowhead. (H and I) Control roots stained on suberin with Fluorol Yellow 088 as (H) a whole root and (I) a cross-section. (J and K) Cu-treated roots stained on suberin with Fluorol Yellow 088 shown as (J) a whole root and (K) a cross-section through area the marked by an arrowhead. (L) Detail of the cross-section from (K). (M) Confocal imaging of the affected apical parts of Cu-treated roots after Fluorol Yellow 088 staining. (N) Three-dimensional reconstruction of phenolic-rich cell walls of the SLAD in Cu- affected roots. Abbreviations: rc, root cap; rh, rhizodermis; cx, cortex; en, endodermis; cc, central cylinder. Scale bars (A, B) = 200 µm; (C–K) = 100 µm; (L, M) = 20 µm. Histochemical analysis and characterization of the root tip deposit Cell walls in stressed tissues are characterized by accumulation of lignin and suberin. We identified the presence of these substances in Cu-treated roots, both in whole roots and in cross-sectionss. Phloroglucinol/HCl staining of the whole root revealed the presence of lignin in a small area in the root tip. On cross-sections, the same staining was localized inside the endodermis and central cylinder (Fig. 2E, F). Staining for suberin with SR 7B also revealed pinkish-red staining in the same part of the root, between xylem vessels and the root tip (Fig. 2G). Similarly, FY088 staining for suberin (Fig. 2H–L) was responsible for strong fluorescence in the same area, visible both in the whole root (Fig. 2J) and in cross-sections (Fig. 2K). On cross-sections, similarly to lignin staining, the highest fluorescent signal originated from the endodermis and central cylinder (Fig. 2L). In control roots, such positive staining was completely absent (Fig. 2H, I). Deposition of phenolics, mainly suberin and lignin, in the endodermis and central cylinder was also revealed by confocal imaging and 3-D reconstruction of the affected area. Phenolics were mainly in cell walls in a short segment of endodermal cells and cells of the central cylinder. These subero-lignified structures in both tissues were localized in an area a few micrometres long, positioned approx. 150–250 µm from the root tip. Affected cells of the endodermis were short, in contrast to elongated cells of the central cylinder. All cells in the central cylinder were characterized by deposition of lignin; none of them showed thickenings typical for xylem vessels or non-uniform deposition of phenolics in cell walls (Fig. 2M, N). Due to the very specific localization in the root apex as well as the dominant occurrence of suberin and lignin in cell walls of this structure we propose to name it a ‘subero-lignified apical deposit’ (SLAD). Measurement of the SLAD position based on the length of endodermal cells along the root axis showed that the SLAD is always present in an area with short, non-elongated endodermal cells, which is characterized by transition between the meristematic and elongation zone. In Cu-treated roots with a formed SLAD, the position of the transition zone is similar to that in control roots, and the length of elongated cells in the elongation zone is lower (Fig. 3). Fig. 3. View largeDownload slide Relationship between cell order from the root initial cells (quiescence centre) and cell length in control (dashed line) and Cu-treated roots (solid line). The grey zone defines cells in treated roots where a SLAD can be formed. Fig. 3. View largeDownload slide Relationship between cell order from the root initial cells (quiescence centre) and cell length in control (dashed line) and Cu-treated roots (solid line). The grey zone defines cells in treated roots where a SLAD can be formed. Evaluation of the effect of locally applied Cu on SLAD formation Roots treated with Cu solely in the root tip region were harvested every 12 h for 2 d and anatomical features were observed. In the control treatment, the root apex did not show any changes in root anatomy, shift of xylem vessels or lateral roots closer to the root apex, or SLAD formation even after 48 h of treatment. However, roots treated with Cu showed symptoms of toxicity. After 12 h of Cu treatment, xylem vessels started to differentiate closer to the root apex but lateral roots or a SLAD were not visible in this zone. After 24 h of Cu treatment, xylem vessels and lateral roots continuously shifted their differentiation and appeared closer to the root apex than in control roots, although a SLAD was still not formed. After 36 h, xylem vessels ceased differentiation and were localized at a very small distance from the root tip. Also, lateral roots appeared closer to the root apex and were visible as small bulges. At this stage, a SLAD was formed as a group of cells stained by FY088. After 48 h, SLAD and xylem vessel differentiation were not different from the previous stage, which indicated cessation of primary root growth, contrary to continuously emerging and growing lateral root primordia (Fig. 4). Fig. 4. View largeDownload slide Schematic representation of the root anatomy influenced by a local application of Cu to a small part of the root tip. In the time course, xylem vessels and lateral roots differentiate closer to the root apex. After 36 h of Cu exposure, SLAD formation occurs. After 48 h, xylem vessels and the SLAD occupy the root tip and stop differentiation, in contrast to continuously growing lateral roots. Fig. 4. View largeDownload slide Schematic representation of the root anatomy influenced by a local application of Cu to a small part of the root tip. In the time course, xylem vessels and lateral roots differentiate closer to the root apex. After 36 h of Cu exposure, SLAD formation occurs. After 48 h, xylem vessels and the SLAD occupy the root tip and stop differentiation, in contrast to continuously growing lateral roots. Analysis of spatial–temporal characteristics of SLAD formation in radish roots The relationship between SLAD formation and Cu exposure time was investigated in a recovery experiment. Ten groups of plants (1–10) were exposed to various durations of Cu 60 treatment and then subsequently transferred to control media for recovery (Table 1). Root morphology changes and SLAD distribution in both primary and lateral roots were calculated. In group 1, i.e. plants growing for the whole experimental period (84 h) in a control solution, almost all lateral roots branched at the basal part of the primary root and the total number of laterals decreased towards the apical part. Both primary and lateral control roots were completely without any SLAD in their apices (Fig. 5A). Plants in groups 2 and 3, exposed to Cu for 1 and 12 h, respectively, had shifted lateral root formation towards the primary root apex. However, the apex of the primary root was completely lacking any SLAD. On the other hand, a SLAD was formed in 9 % of lateral roots after 1 h exposure and in 20 % after 12 h exposure. In both groups, all SLADs forming laterals were localized at the basal part of the primary root (Fig. 5B). Moreover, some lateral roots showed a region close to the apex, where xylem vessels were short and stacked but without a formed SLAD (data not shown). Because this region was followed by elongated xylem vessels, it possibly indicates a partial decrease in root growth during Cu exposure followed by new growth without SLAD formation. Plants in groups 4–9, exposed to Cu from 24 to 84 h, showed typical morphological changes induced by metal toxicity. In contrast to groups 2 and 3, the primary roots in groups 4–9 always formed a SLAD in their apices and the distribution of lateral roots was more shifted towards their tips. In groups 4–9, a total of 34–53 % of lateral roots formed a SLAD in their apices. The formation of a SLAD was the most prominent in those lateral roots that were developed close to the primary root base and decreased towards the primary root apex. The laterals branched at the apical part of the primary root did not form any SLAD (Fig. 5C). Group 10, fully exposed to Cu for 168 h without any recovery, showed a SLAD in the primary root apex, and nearly 70 % of all lateral roots also formed a SLAD (Fig. 5D). Moreover, deposition of suberin into SLADs in lateral roots also affected other endodermal cells and was visible as a differentiated endodermis along the whole lateral root axis (Fig. 5E). Fig. 5. View largeDownload slide Root morphological changes and distribution of SLAD-affected roots (red root) and roots without a SLAD (black root) after the recovery experiment. Percentages show the number of SLAD-affected lateral roots in individual groups. Root lengths on the schemes are illustrative, but the distribution and proportion of lateral roots with or without a SLAD depict the real situation. (A) Scheme of a control root (group 1), fully exposed to control solution, that does not form a SLAD in laterals or the main root. (B) Scheme of morphological changes of a root after 12 h of Cu exposure followed by a 72 h recovery time (group 3). Note the lack of SLAD formation in the primary root, shifted lateral root production towards the root apex and formation of a SLAD in 20 % of lateral roots. (C) Scheme of morphological changes of a root after 72 h of Cu exposure followed by a 12 h recovery time (group 8). A SLAD is formed in the primary root and in 45 % of all lateral roots. (D) Scheme of root morphology after a 168 h exposure to Cu (group 10). Around 70 % of all lateral roots form a SLAD, and a SLAD is also present in the primary root. (E) A lateral root (LR) after a 168 h exposure to Cu (group 10) and FY088 staining. The presence of suberin is visible in the SLAD (arrowhead) and in all endodermal cells (asterisk) from the SLAD to the junction on the primary root (PR). Fig. 5. View largeDownload slide Root morphological changes and distribution of SLAD-affected roots (red root) and roots without a SLAD (black root) after the recovery experiment. Percentages show the number of SLAD-affected lateral roots in individual groups. Root lengths on the schemes are illustrative, but the distribution and proportion of lateral roots with or without a SLAD depict the real situation. (A) Scheme of a control root (group 1), fully exposed to control solution, that does not form a SLAD in laterals or the main root. (B) Scheme of morphological changes of a root after 12 h of Cu exposure followed by a 72 h recovery time (group 3). Note the lack of SLAD formation in the primary root, shifted lateral root production towards the root apex and formation of a SLAD in 20 % of lateral roots. (C) Scheme of morphological changes of a root after 72 h of Cu exposure followed by a 12 h recovery time (group 8). A SLAD is formed in the primary root and in 45 % of all lateral roots. (D) Scheme of root morphology after a 168 h exposure to Cu (group 10). Around 70 % of all lateral roots form a SLAD, and a SLAD is also present in the primary root. (E) A lateral root (LR) after a 168 h exposure to Cu (group 10) and FY088 staining. The presence of suberin is visible in the SLAD (arrowhead) and in all endodermal cells (asterisk) from the SLAD to the junction on the primary root (PR). DISCUSSION There is knowledge about development of root hairs or lateral roots very close to the apex in the case of a stress-induced morphological response (Potters et al., 2007). Another stress-induced plant response is production and deposition of phenolic substances, e.g. suberin or lignin, into damaged cells as well as enhancement of development of root apoplasmic barriers when plants suffer under hostile conditions (Caño-Delgado et al., 2003; Enstone et al., 2003). Similar symptoms were observed in radish plants exposed to a high concentration of Cu in our experiments. Moreover, we found specific deposition of the phenolics suberin and lignin inside a root tip area that we described as a subero-lignified apical deposit (SLAD). This is in contrast to the well accepted model of root endodermis suberization which is based on continuous differentiation of cells from the root base to the apex. In the case of endodermal suberization, it is not only an environmentally, but also a positionally regulated process. Unilateral exposure of maize roots to a high concentration of Cd caused decreased cell elongation activity in the Cd-exposed part of the root and, simultaneously, cells of this part deposited suberin in cell walls more intensively and closer to the root tip in contrast to the Cd-free root site (Lux et al., 2011). Endodermal cells positioned near the phloem poles start to suberize much closer to the root tip than other endodermal cells, and the number of suberized cells increases toward the base. Suberization of the endodermis is not completed until all endodermal cells in the basal part of the root are completely covered by suberin. The exception is a group of passage cells, located near to the xylem poles that suberize only occasionally, as was shown in garlic (Allium cepa L.) (Waduwara et al., 2008) and soybean [Glycine max (L.) Merr.] roots (Thomas et al., 2007). Previous observations on A. thaliana showed that emerging lateral roots induced suberization in a well-defined group of endodermal cells located above the lateral root primordia or around emerged lateral roots. Such cells may suberize prematurely, very early and during the first stages of lateral root growth, and may protect primary root from an uncontrolled influx of solutes (Martinka et al., 2012). In SLADs, deposition of phenolics into endodermal cells can be separated from the rest of the endodermis. As revealed by the experiment with Cu stress applied solely to the root tip, SLADs can be formed independently and endodermal cells can suberize prematurely in this part of the root, compared with other endodermal cells. Premature suberization of endodermal cells in the endodermal part of SLADs is peculiar for several reasons. According to cell length measurement, a SLAD affects cells in the basal part of the meristematic zone. Normally, young cells in apical meristems are undifferentiated and, for example, in A. thaliana roots, local ablation of an atrichoblast cell in the rhizodermis induces changes in a trichoblast cell located nearby which does not develop into a root hair but takes the position and function of an ablated atrichoblast cell (Berger et al., 1998). The fate of cells in the meristematic zone is not strongly pre-determined. If all meristematic cells are exposed to the same conditions and environment, but only endodermal and central cylinder cells start to produce secondary metabolites such as phenolics, they are probably pre-determined to have such a function very early. In the case of the endodermis, such a reaction is supported by the important position of this layer on the border between internal vascular tissues and the external environment (Robbins et al., 2014). Current knowledge about the very high plasticity of endodermis development and suberin deposition influenced by many signals and environmental conditions can also support these findings (Barberon, 2017). Even older data showed the presence of phenolic-storing cells in plant tissues which react very rapidly to stress and subsequently may become lignified or suberized (e.g. Beckman, 2000, and references therein). One layer characteristic of such cells is the endodermis (Mueller and Beckman, 1976). Pro-endodermis, a layer of young endodermal cells before their differentiation, is in some species also characterized by a higher phenolic content (Beckman, 2000; Ma and Peterson, 2003). Auxin hormonal regulation can also explain early endodermis differentiation during SLAD formation. Auxin transported from the root base to the apex changes direction at the root apex, and local gradients of this hormone induce maintenance of meristematic activity and various reactions and developmental responses (Moubayidin et al., 2009; Overvoorde et al., 2010). If unfavourable conditions change the hormonal concentrations [through degradation of reactive oxygen species or peroxidases by changes of pH, failure in transport or incorrect localization of the cytoskeleton (Pasternak et al., 2005)], the sensitive root tip may react very rapidly and reactions start preferentially in these areas. Deposition of phenolics into the narrow area of the central cylinder encircled by differentiated endodermal cells can be related to differentiation of xylem, even though vessels are not visible in this part of the SLAD. Normally xylem differentiation induces deposition of lignin and cell death in elongated cells in the central cylinder (Srivastava and Singh, 1972; Fukuda, 1997; van Doorn and Woltering, 2005). If initial cells in meristems of A. thaliana roots are under DNA-damaging stress (γ-rays, UV light, heavy metals, reactive oxygen species or toxic compounds), they are selectively eliminated and killed. Because initials produce lineages of cells and genetic information, damage induced by stress can lead to mutation in daughter cells, too (Fulcher and Sablowski, 2009). Therefore, DNA-damaging factors in initials leads to programmed cell death, and dead cells are in the meristematic apical zone observable after propidium iodide staining as intensively stained structures (Furukawa et al., 2010). Also, despite common high susceptibility of all initials to DNA damage, cells of the central cylinder are much more sensitive to damage and cell death (Fulcher and Sablowski, 2009). Therefore, higher susceptibility and possible cell death of cells in the central cylinder might relate to the observed stress-induced deposition of phenolics in our experiments. Radish root morphology affected by Cu, including an increased number of lateral roots, their shift to the root apex and modified deposition of phenolics, is in agreement with Cu-induced reorganization of A. thaliana root architecture (Lequeux et al., 2010). The described morphological changes in roots are similar to stress reactions called stress-induced morphogenic responses (SIMRs) (Potters et al., 2007). According to these authors, in the case of an SIMR, stress does not kill the plant, but causes (1) inhibition of elongation; (2) local stimulation of cell division; and (3) alterations to cell differentiation. Subsequently, a new developmental programme completely changes the morphology of the stressed plant (Potters et al., 2009). Due to this, we assume that the formation of SLADs can be one part of the SIMR reaction, because (1) SLAD-affected roots ceased their elongation activity; (2) SLAD formation occurs in roots along with apically stimulated lateral root growth; and (3) differentiation of cells in SLADs is altered by deposition of suberin and lignin. According to the recovery experiment, 1 h of Cu 60 exposure is enough to induce SLAD formation in some lateral roots, and exposure for >24 h induces SLAD formation in half of all lateral roots regardless of the recovery time. However, if SLAD formation is a part of the SIMR, deposition of phenolics into SLADs is very probably not a reason for but a reaction to limited growth of the root. Because SLAD formation follows and does not precede emergence of laterals in the apex, it is also not responsible for reorganization or remodelling of the root system. It is similar to the reaction of wheat roots to Al stress, when lignification affects sensitive roots. Deposition of phenolics into elongating cells is not a reason for cessation of growth, but a reaction to damage caused by the element in excess (Sasaki et al. 1996). In the case of the endodermis, SLAD-forming deposits are typically located in the cell walls of short cells present just before the onset of elongation. The change from meristematic to elongation activity occurs normally in the transition zone. The transition zone is composed of developmentally plastic cells (Verbelen et al., 2006), and plasticity allows the root to undergoe diverse tropisms and react to internal or external stimuli (Baluška et al., 2010). Changes in root growth in the transition zone are affected by reaction to nitrate (Trevisan et al., 2015), osmotic stress (Baluška and Mancuso, 2013) or Al stress (Sivaguru and Horst, 1998), among others. Cells of the transition zone are rich in photosynthates transported by phloem differentiating just in this portion of the root (Verbelen et al., 2006). The transition zone is also responsible for root growth regulation via phytohormones (Dello Ioio et al., 2007; Ubeda-Tomás et al., 2009). High sensitivity, unloading of metabolites and high oxygen influx despite low meristematic activity in the transition zone (Baluška and Mancuso, 2013) provide suitable conditions for deposition of phenolics and SLAD formation in this zone. The role of the SLAD and deposition of phenolic-containing substances into a narrow area of the transition zone between xylem vessels and root initials in the SIMR reaction may be in exclusion of supplies of sugar and metabolites to the quiescence centre or initial cells to restrict growth in the case of unfavourable conditions. Similarly, the SLAD may prevent xylem vessels from taking up unfavourable solutes. An increased number of lateral roots and their shift to the root apex in the SIMR reaction can compensate for loss of primary root activity but, if unfavourable conditions continue, all lateral roots stop growth and form a SLAD. Moreover, in such negative conditions, phenolics (suberin) are present not only in SLADs, but in all endodermal cells from the root base to the SLAD. Such an isolation of the root from the negative environment by phenolics deposited into SLADs and the endodermis may also explain the formation and existence of SLADs in the root apex. Conclusion Our recent findings revealed the presence of a specific structure formed in the root apex of radish after Cu exposure. This structure is localized in the transition zone of the root and consists of phenolic-enriched components lignin and suberin. Therefore, we named this structure ‘subero-lignified apical deposit’ or SLAD. Formation of a SLAD depends on the time of exposure to high Cu concentrations and affects endodermal and central cylinder cells in both primary and lateral roots. Moreover, it is connected to stress-induced remodelling of the root system, such as a decrease in root growth or an increase in lateral root production. The exact role of a SLAD can only be predicted at present; however, we assume that its presence in the root tip may enhance endodermal barrier function, decrease xylem conductivity or limit phloem transport. Future investigations with different kinds of stresses and with various plant species can answer this question and determine whether a SLAD is a more general phenomenon. ACKNOWLEDGEMENTS The work was supported by the Slovak Grant Agency VEGA grant VEGA 1/0605/17. The authors thank members of the Core Facility CIUS of Vienna University for the possibility to work with the confocal laser scanning microscope and their help and technical assistance. They also thank all reviewers and the Handling Editor, Dr Nigel Chaffey, for comments, improvements and suggestions provided during manuscript revision. LITERATURE CITED Andersen TG, Barberon M, Geldner N. 2015. Suberization—the second life of an endodermal cell. Current Opinion in Plant Biology  28: 9– 15. Google Scholar CrossRef Search ADS PubMed  Baluška F, Mancuso S. 2013. Root apex transition zone as oscillatory zone. Frontiers in Plant Science  4: 1– 15. 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Published: Feb 10, 2018

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