JXB Advance Access originally published online on November 3, 2006
Journal of Experimental Botany 2006 57(15):4201-4213; doi:10.1093/jxb/erl197
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RESEARCH PAPER |
Aluminium toxicity in plants: internalization of aluminium into cells of the transition zone in Arabidopsis root apices related to changes in plasma membrane potential, endosomal behaviour, and nitric oxide production
1ek Baluka1,2
ka1,*
1Institute of Botany, Slovak Academy of Sciences, Dubravska cesta 14, SK-845 23, Bratislava, Slovakia
2Institute of Cellular and Molecular Botany, University of Bonn, Bonn, Germany
3Institution of Cell Imaging and Ultrastructure Research, University of Vienna, Vienna, Austria
* To whom correspondence should be addressed. E-mail: miroslav.ovecka{at}savba.sk
Received 6 June 2006; Accepted 11 September 2006
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Abstract |
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The extent of aluminium internalization during the recovery from aluminium stress in living roots of Arabidopsis thaliana was studied by non-invasive in vivo microscopy in real time. Aluminium exposure caused rapid depolarization of the plasma membrane. The extent of depolarization depends on the developmental state of the root cells; it was much more extensive in cells of the distal than in the proximal portion of the transition zone. Also full recovery of the membrane potential after removal of external aluminium was slower in cells of the distal transition zone than of its proximal part. Using morin, a vital marker dye for aluminium, and FM4-64, a marker for endosomal/vacuolar membranes, an extensive aluminium internalization was recorded during the recovery phase into endosomal/vacuolar compartments in the most aluminium-sensitive cells. Interestingly, aluminium interfered with FM4-64 internalization and inhibited the formation of brefeldin A-induced compartments in these cells. By contrast, there was no detectable uptake of aluminium into cells of the proximal part of the transition zone and the whole elongation region. Moreover, cells of the distal portion of the transition zone emitted large amounts of nitric oxide (NO) and this was blocked by aluminium treatment. These data suggest that aluminium internalization is related to the most sensitive status of the distal portion of the transition zone towards aluminium. Aluminium in these root cells has impact on endosomes and NO production.
Key words: Aluminium internalization, Arabidopsis thaliana, endosomal compartments, live cell microscopy, membrane potential, morin vital staining, nitric oxide, recovery, root transition zone, vacuoles
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Introduction |
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Aluminium toxicity is an important growth-limiting factor in acid soils. The main symptom of aluminium toxicity is the dramatic inhibition of root growth. Some decades ago, two pioneer works postulated that the decreased root growth is a consequence of the inhibition of cell division (Clarkson, 1965) and cell elongation (Klimashevski and Dedov, 1975). Later Ryan et al. (1993) recognized the root apex as a primary site of aluminium-induced injury in plants. More recently, numerous reports in the literature describe the aluminium-induced changes occurring particularly in the apical regions of the root, leading to expression of aluminium-toxicity symptoms: changes in root cell patterning (Doncheva et al., 2005), irregular cell division, alterations in cell shape, and vacuolization (Vázquez et al., 1999;
iamporová, 2000), cell wall thickening and callose deposition (Horst et al., 1999; Nagy et al., 2004; Jones et al., 2006), disintegration of the cytoskeleton (Sivaguru et al., 1999, 2003b), formation of myelin figures and membranous electron-dense deposits (Vázquez, 2002), disturbance of plasma membrane properties (Miyasaka et al., 1989; Olivetti et al., 1995; Pavlovkin and Mistrík, 1999; Sivaguru et al., 1999, 2003b; Ahn et al., 2001, 2002, 2004; Ahn and Matsumoto, 2006), as well as the production of reactive oxygen species (Darkó et al., 2004; Jones et al., 2006). These reactions are only few examples of how aluminium affects the root cells.
The root apex consists of the zone of cell division (meristem), followed by the distal and proximal transition zone where cells are prepared for rapid cell expansion in the elongation zone (Baluka et al., 1990, 1994, 1996; Ishikawa and Evans, 1993; Verbelen et al., 2006). Sivaguru and Horst (1998) discovered that the distal part of the transition zone (DTZ) is the most sensitive part of the root to aluminium stress. Using a sophisticated experimental approach, they revealed that local applications of aluminium to the rapidly elongating cells do not inhibit their growth, while local applications of aluminium to cells of the distal portion of the transition zone dramatically inhibit root growth. However, it remains elusive which processes specific to this small developmental window in root cell development are particularly sensitive to aluminium. Studies by Kollmeier et al. (2000) indicate that the unique status of auxin in cells of the distal portion of the transition zone could be responsible for this high sensitivity of DTZ cells to aluminium.
One of the most relevant problems of recent research on aluminium phytotoxicity is to define the primary site of its action at a cellular and subcellular level. The crucial question is whether aluminium acts primarily in the apoplast or in the symplast. Consequently, the determination of primary cellular mechanisms responsible for the rapid cell response to the aluminium toxicity is still a matter of discussion. Therefore, it is necessary to characterize the uptake of aluminium into the root cells and to monitor its spatial and temporal distribution in cells of living roots.
Vital staining is one of the best and rapid methods for monitoring aluminium localization and distribution in plants. At present, despite some negative views (Eticha et al., 2005), the aluminium-specific fluorescent dye morin as well as lumogallion seem to be the best vital fluorescent dyes for aluminium detection. Both of them appear effective in the nanomolar range of aluminium concentrations. The morin-staining method showed that aluminium was rapidly taken up into cultured tobacco BY-2 cells (Vitorello and Haug, 1996), wheat (Tice et al., 1992), and maize root cells (Jones et al., 2006). Lumogallion staining confirmed the rapid aluminium accumulation in soybean root cells (Silva et al., 2000). On the other hand, Ahn et al. (2002) reported that most of the aluminium detected by morin was located preferentially in the cell walls of squash root cells within the first 3 h of an experiment. Transmission electron microscopy studies in combination with energy-dispersive X-ray analysis are approaches giving more detailed information about the distribution of aluminium at the subcellular and ultrastructural level. Vázquez et al. (1999) observed the presence of aluminium in cell walls and vacuoles of maize root tip cells after 4 h of aluminium treatment. Interestingly, 24 h of exposure resulted in abundant occurrence of aluminium deposits within vacuoles, but less in cell walls. Accelerator mass spectrometry in single cells of Chara corallina revealed the uptake of aluminium into the cytoplasm during the first 30 min followed by its sequestration into vacuoles, although intracellular aluminium represented only 0.5%; the major portion being apoplastic (Taylor et al., 2000).
Despite extensive research efforts focusing on aluminium uptake, results are often conflicting. Most of the discrepancies arise from different methods and experimental conditions (concentrations of aluminium, sensitivity of methods, exposure times, cell types, sample processing, etc). In the studies on intact roots, the particular stage of cellular development (Baluka et al., 1996), reflecting its different sensitivity to aluminium (Sivaguru and Horst, 1998; Sivaguru et al., 1999), was not always addressed with respect to aluminium internalization. Therefore, it is difficult to generalize our knowledge about the uptake of aluminium into plant cells. Moreover, data on the fate of aluminium internalized during plant recovery are completely missing.
To gain a synoptic understanding of aluminium effects, the extent of aluminium internalization into well-defined cells of living roots of Arabidopsis thaliana was studied in a continuous mode under controlled conditions by non-invasive live microscopy. It is reported that those cells which are most aluminium-sensitive are also the most active in the internalization of apoplastic aluminium during recovery. Importantly, endosomal and vacuolar compartments are highly enriched with the internalized aluminium only in cells of the distal portion of the transition zone.
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Materials and methods |
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Plant material and growth conditions
Seeds of Arabidopsis thaliana L., ecotype Columbia, were surface-sterilized with 0.25% sodium hypochlorite for 3 min, washed and sown on an agar-solidified nutrient medium in Petri dishes. The nutrient medium was based on Murashige-Skoog salts (Murashige and Skoog, 1962) with addition of vitamins (myo-inositol 10 mg l–1, calcium pantothenate 0.1 mg l–1, niacin 0.1 mg l–1, pyridoxin 0.1 mg l–1, thiamin 0.1 mg l–1, biotin 0.001 mg l–1 of medium), FeSO4.7H2O (1.115 mg l–1), CaCl2 (111 mg l–1), sucrose (10 g l–1), and agar (10 g l–1), the final pH was adjusted to 4.5. The seeds were vernalized at 4 °C for 24 h. Petri dishes were placed into a growth chamber, positioned vertically and kept under controlled environmental conditions at 25 °C, 180 µmol m–2 s–1 and a 12/12 h day/night rhythm.
Root growth and detection of aluminium
Effects of aluminium on root growth were observed on seedlings which, 2 d after germination (DAG), were transferred to Petri dishes containing agar-solidified nutrient medium with different aluminium concentrations (0, 10, 50, 100, 200, and 300 µM total concentration of Al in the form of AlCl3.6H2O). Root elongation was measured every day during a 7-d period. After 7 d of cultivation, aluminium was detected by staining whole roots with haematoxylin (Polle et al., 1978) or morin (Vitorello and Haug, 1997). Roots stained with haematoxylin were observed in bright field and morin fluorescence was visualized by an Olympus BX51 microscope (Olympus, Japan) equipped with BP 470–490 excitation filter, BA 515 IF barrier filter and a DM 505 dichroic mirror.
Electrophysiology
Measurements of the plasma membrane electrical potential difference (Em) were carried out at 24 °C in root cells of intact Arabidopsis seedlings, 2 DAG, by the standard microelectrode technique as described by Pavlovkin et al. (1993). The perfusion solution contained 0.1 mM KCl, 1 mM Ca(NO3)2, 1 mM NaH2PO4, 0.5 mM MgSO4; pH was adjusted to 4.5. Diffusion potential (ED) was determined by application of inhibitors (1 mM NaCN+1 mM SHAM) dissolved in perfusion solution. The influence of morin on Em was measured upon adding 100 µM morin to the perfusion solution. Effects of aluminium on Em were monitored during continual exchange of the perfusion solution by the experimental solution (50 µM AlCl3) in perfusion solution, perfusion speed 5 ml min–1). After 5 min or 30 min of aluminium exposure the experimental solution was washed out with perfusion solution. Changes of Em induced by aluminium were measured continuously during the whole experiment. Insertion of the microelectrode into the cortical cells of the distal portion of the transition zone, located 150–300 µm behind the root tip, and of the proximal portion of the transition zone, located 300–400 µm behind the root tip (Verbelen et al., 2006), was performed under microscopic control. Measurements were evaluated separately for each zone.
Live microscopy of aluminium internalization into the cells
Arabidopsis seedlings 2 DAG were transferred to microscopic slides modified to micro-chambers by coverslips (Oveka et al., 2005). The chambers were filled with liquid nutrient medium of the same composition as used for cultivation in Petri dishes, but without agar and placed into sterile glass cuvettes containing the same nutrient medium (pH 4.5). The seedlings were grown in a vertical position under light in the growth chamber. During a 12-h period the seedlings resumed stable root growth. Subsequently the micro-chambers were gently perfused with the aluminium-containing medium (50 µM AlCl3 in nutrient solution, pH 4.5, perfusion speed 10 µl min–1). After a 30 min pulse treatment, aluminium was washed out and the roots of the Arabidopsis seedlings were stained with 100 µM morin for 20 min, using the same perfusion technique and then washed with the nutrient solution. After this exchange, internalization experiments were performed in the micro-chambers directly on the microscope stage during 3 h. For labelling endosomes and tonoplast, the styryl dye FM4-64 (N-(3-triethylammoniumpropyl)-4-(8-(4-(diethylamino) phenyl)hexatrienyl)pyridinium dibromide) was used at a final concentration of 4 µM in nutrient solution, applied for 5 min prior to the aluminium treatment. Internalization of aluminium was observed in living roots in real time by an inverted microscope Leica DM IRE2, equipped with the confocal laser scanning system Leica TCS SP2 (Leica Microsystems Heidelberg, Germany). Excitation wavelength for FM4-64 was 514 nm and for morin 458 nm. Fluorescence was observed between 640 nm and 700 nm (FM4-64) and 480 nm and 510 nm (morin).
FM4-64 internalization and BFA-induced compartment formation
Working solution of 5 µM FM4-64 was prepared from the stock solution (1 mg ml–1 FM4-64 in DMSO). With FM4-64 the plants were incubated for 10 min and the dye was washed out before observation. Roots labelled for 5 min with the FM 4-64 were pretreated with BFA applied at a concentration of 35 µM for 25 min. The effect of aluminium was studied by the application of 90 µM AlCl3 for 90 min before treatment with BFA and FM4-64.
NO labelling and measurements in control and aluminium-treated root cells
Detection of nitric oxide (NO) was performed by the specific fluorescent probe 4,5-diaminofluorescein diacetate (DAF-2 DA; Calbiochem, USA). The roots were incubated with 15 µM DAF-2 DA for 30 min and washed before observation. As a negative control, they were treated with 10 µM of the NO-scavenger cPTIO (Lombardo et al., 2006). Examination was performed with a confocal laser scanning microscope system using standard filters and collection modalities for DAF-2 green fluorescence (excitation 495 nm; emission 515 nm). Fluorescence intensity was measured with the open source software Image-J (http://rsb.info.nih.gov/ij/).
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Results |
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Influence of aluminium on root elongation and morphology
Arabidopsis seedlings cultivated for 7 d on agar plates with different concentrations of AlCl3 exhibited concentration-dependent inhibition of root growth (Fig. 1). Growth of the primary roots was only slightly reduced by 10 µM AlCl3 (to 95% of the control values, not statistically significant according to t test at P=0.05). Inhibition of root growth was statistically significant at 50 µM AlCl3 as compared to control according to t test at P=0.01. However, growth of the primary roots was reduced only to 89% of the control. The inhibition was more apparent at 100 µM and 200 µM AlCl3 (59% and 45% of control growth, respectively, highly significant at P=0.001), while root growth was fully inhibited by 300 µM AlCl3 (only 2% root elongation as compared to control plants, highly significant at P=0.001; Figs 1, 2). Severe inhibition of root elongation was accompanied by radial expansion of the root cells at 100 µM and 200 µM AlCl3, but not at 300 µM AlCl3, due to the complete inhibition of root growth (Fig. 3). Root architecture of Arabidopsis seedlings exposed to lower concentrations of aluminium (10–100 µM AlCl3) was almost unaffected. However, 200 µM and 300 µM AlCl3 induced enhanced formation of lateral roots (Fig. 2). Thus, the inhibition of root elongation was tightly correlated with, and partly compensated by, an increasing number of growing zones with the formation of lateral roots.
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Histochemical detection of aluminium
For histochemical detection of aluminium in the roots of Arabidopsis plants cultivated on agar plates with different concentrations of aluminium, two different staining methods were used: haematoxylin staining and morin staining (Polle et al., 1978; Vitorello and Haug, 1997). Using both methods, an aluminium-specific signal was observed mainly in the apical parts of the roots exposed to aluminium for 7 d (Fig. 3). In the control (Fig. 3), and at 10 µM AlCl3 (data not shown), aluminium staining by haematoxylin was not detectable and only a weak diffuse fluorescence of morin occurred. At 50 µM AlCl3, there was hardly any detection of aluminium in roots by means of haematoxylin, while morin gave bright fluorescence (Fig. 3). At higher concentrations, haematoxylin started to detect aluminium in the roots, the pattern of staining being similar to the morin fluorescence. In the roots grown at 100 µM and 200 µM AlCl3, both staining methods showed maximum accumulation of aluminium in the root apex. In the plants exposed to 300 µM AlCl3, aluminium was abundant not only in the root apex but it also invaded the root central cylinder (Fig. 3).
Obviously, the sensitivity of morin to aluminium is higher than that of haematoxylin. Thus, morin is considered as a more suitable tracer dye for aluminium detection in the cells of the Arabidopsis root apex grown on aluminium-supplemented agar plates. The critical discrimination limit in the sensitivity between morin and haematoxylin is apparently at 50 µM AlCl3 (Fig. 3). Because of mild effects on root growth and the capability of morin to localize aluminium, 50 µM AlCl3 was utilized as the indicative testing concentration for monitoring aluminium effects on Arabidopsis roots in the following experiments.
Effects of aluminium on the plasma membrane electrical potential
In order to detect immediate cell responses of the root apex to aluminium, the plasma membrane potential (Em) was recorded before and during aluminium application, as well as after the removal of aluminium by washing. Em in the cells of the root cortex revealed distinct properties in different developmental zones. In this case, the distal part of the transition zone, just behind the cell division zone, was located 150–300 µm behind the root tip, the proximal part of the transition zone 300–400 µm behind the root tip (Verbelen et al., 2006). Em of cortical cells varied between –82 mV and –98 mV (–88±4 mV, n=37) in the distal part of the transition zone (DTZ) and between –94 mV and –117 mV (–105±7 mV, n=29) in the proximal part (PTZ). Based on this electrophysiological characteristic, all further experiments were performed separately for DTZ and PTZ.
The diffusion potential (ED) was determined in order to distinguish between passive and active, i.e. energy-dependent, components of the Em by application of inhibitors (1 mM NaCN+1 mM SHAM); the Em of cortical cells rapidly depolarized in both DTZ and PTZ to ED (–40 to –41 mV, Fig. 4). As a control, 100 µM morin revealed almost no detectable changes of Em (data not shown); thus, morin does not significantly affect the plasma membrane properties. This confirms the suitability of this dye for studying aluminium distribution in living cells.
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The main objective of the present electrophysiological experiments was to characterize dynamic changes of Em and to determine the sensitivity of the two developmental zones (DTZ and PTZ) during the exposure to 50 µM AlCl3. Aluminium-induced depolarization of Em occurred within 2 min after aluminium application in both developmental zones (Fig. 5A). However, the membrane potential depolarized further in the cells of DTZ (to ED) than in the cells of PTZ (–78 mV to –88 mV). Complete repolarization of Em was achieved by removing aluminium from the perfusion solution within 10 min in the cells of DTZ, while the cells of PTZ repolarized within only 3 min (Fig. 5A). While the period of treatment used in the aluminium internalization experiments was 30 min, the effects of short-term aluminium treatment (5 min; Fig. 5A) were compared with the 30 min exposure (Fig. 5B). Extent and speed of the Em depolarization were similar (data not shown), but the speed of repolarization was again different in the two developmental zones. After aluminium removal from the solution, the time required for complete repolarization was 14 min in the cells of DTZ and only 6 min in the cells of PTZ (Fig. 5B). Apparently DTZ is much more sensitive to aluminium than PTZ. Nevertheless, depolarization of the plasma membrane was fully reversible, which was consequently followed by recovery of root growth (data not shown).
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Internalization and accumulation of aluminium within endosomal/vacuolar compartments
The dynamics of aluminium internalization was monitored in the root cells of Arabidopsis seedlings 2 d after germination. The intact roots were treated with 50 µM AlCl3 for 30 min, washed and stained with morin. After washing out morin, the seedlings were kept in control medium for recovering and the time-course of aluminium internalization in the living root apices could be observed with a confocal microscope for 3 h. In cells of meristem and DTZ, aluminium was located exclusively in the apoplast during the first 20 min of recovery (after aluminium removal; Fig. 6A). From then on, it was internalized into the cells. A first detectable diffuse fluorescence signal of morin-stained aluminium in the cytoplasm appeared after 60 min (Fig. 6B). Aluminium further proceeded into roundish structures with blurred edges, and 2 h and 30 min after the end of treatment it was located in vacuole-like structures of different size with clearly defined boundaries. In the cytoplasm the signal became weaker, in the cell walls it remained present (Fig. 6C). After 3 h and 30 min, these cells accumulated aluminium almost exclusively in the vacuole-like compartments (Fig. 6D).
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The assumption that the target compartment of aluminium sequestration in the cells is the vacuole was confirmed by FM4-64, the dye widely used for labelling the plasma membrane, endocytic membranous compartments and tonoplast (Betz et al., 1996; Geldner, 2004; Ove
ka et al., 2005; 
amaj et al., 2005). Accumulation of aluminium was shown in the cells of root developmental zones that, as evident from the previous experiments, revealed different sensitivity to aluminium (Fig. 5). In the meristematic cells (Fig. 7A) and in the cells of the distal portion of the transition zone (Fig. 7B), aluminium was internalized rapidly and accumulated into the vacuolar compartments within 2 h and 50 min of recovery. Tonoplast was labelled red with FM4-64 whereas the aluminium-containing lumen of the vacuoles was stained green with morin (Fig. 7A, B).
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In the cells of the proximal portion of the transition zone, there was no prominent accumulation of aluminium even 3 h 10 min after aluminium deprivation (Fig. 7C). Moreover, aluminium did not enter vacuoles neither in the proximal portion of the transition zone nor in the elongation zone, it could only be detected in the apoplast. This indicates that after the removal of free aluminium ions from the medium, residual aluminium bound to the cell wall can be internalized into endosomal compartments and vacuoles of living cells of the meristem as well as of the distal portion of the transition zone. On the contrary, in the less sensitive cells of the proximal portion of the transition zone, as well as in the elongation region (data not shown), cell wall-bound aluminium is not internalized and remains located in the apoplast.
Effects of aluminium on endosomal behaviour
After 10 min exposure of roots, FM4-64 strongly labelled cross-walls of the cells as well as the early endosomes (Fig. 8A; Voigt et al., 2005). When such cells were exposed to brefeldin A (BFA), the early endosomes aggregated into the so-called BFA-induced compartments (Fig. 8B; amaj et al., 2004, 2005). Aluminium treatment prevented formation of such BFA-induced compartments (Fig. 8C, D), suggesting that early endosomes, involved in aluminium internalization, were affected.
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Effects of aluminium on NO production
NO-specific DAF-2DA labelling showed spatial distribution of NO production in cells of control root tips (Fig. 9A) and local changes of this distribution induced by aluminium treatment (Fig. 9B). After application of the NO scavenger cPTIO, the DAF-2DA fluorescence signal was lacking, indicating effective NO scavenging (Fig. 9C). In control root apices, there were three local centres of NO production: one at the root cap statocytes, another one at the quiescent centre and distal portion of the meristem, and the third, the most prominent one, at the distal part of the transition zone (the blue line in Fig. 9D). While the NO-scavenger stopped NO production in roots (the green line in Fig. 9D), aluminium treatment (90 µM for 1 h) completely abolished only the NO production peak in the distal part of the transition zone; the first two peaks became even more pronounced (the red line in Fig. 9D).
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Discussion |
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Although aluminium toxicity in plants has been extensively studied from different points of view, a complete image of its distribution at the cellular level is still missing. In line with the supporting data for aluminium uptake into the cells, evidence for predominant accumulation of aluminium only in the apoplast has also been given. However, it must be kept in mind that much of the data available in this field were obtained with different plant species and under experimental conditions which were not comparable. Experimental approaches such as the detection of aluminium in living cells with high sensitivity in physiological conditions may contribute to clarifying this situation.
Internalization and distribution of aluminium can be visualized at low concentrations in living cells
The time-course of internalization of aluminium in actively growing root cells of Arabidopsis thaliana was detected by the application of non-invasive microscopy techniques. Both dyes morin and FM4-64 were used as vital markers in living cells under microscopic control. While FM4-64 is widely used as a vital marker of endocytosis in different cell types (Vida and Emr, 1995; Betz et al., 1996; Fischer-Parton et al., 2000; Bolte et al., 2004; Oveka et al., 2005; amaj et al., 2005; Dettmer et al., 2006; Dhonukshe et al., 2006), morin was mainly used as a last staining step in the final stage of the experiments. In this study, morin was used as a marker of aluminium redistribution in living Arabidopsis root cells. Morin labelling in the early stages of recovery and careful visualization of its fluorescence allowed the time-course of aluminium internalization to be studied at low, non-lethal concentrations even in the most sensitive cells of DTZ.
Cells of various root developmental zones have different sensitivity to aluminium and show specific patterns of recovery
The effect of aluminium on root cells became evident by rapid changes of the electrical membrane potential, which were different in the cells of various developmental stages. The root apex consists of distinct developmental zones including the cell division zone (meristem), two zones of preparation for rapid cell expansion (DTZ and PTZ), followed by the actual zone of rapid cell elongation (Baluka et al., 1990, 1994, 1996; Ishikawa and Evans, 1993; Verbelen et al., 2006). Aluminium caused the rapid depolarization of the plasma membrane electro-potential (Em) in the cells of both the DTZ and PTZ. The extent of depolarization, however, was much greater in the more sensitive DTZ. This is in accordance with the observations by Sivaguru and Horst (1998), Horst et al. (1999), and Sivaguru et al. (1999, 2003a), who described a different sensitivity of the cells in different developmental zones. This clearly indicates that the extent of the aluminium sensitivity as a function of cellular developmental stages should be taken into consideration.
Concerning recovery from aluminium stress, removing free aluminium from the medium was followed by full regeneration of the Em values. Hence, the changes in electrophysiological properties of the plasma membrane induced by aluminium were reversible under the experimental conditions in the recovering cells of both DTZ and PTZ. Consistent with developmentally dependent differences in sensing aluminium, the process of plasma membrane recovery was slower in the cells of the DTZ as compared to those of PTZ.
The cellular distribution of aluminium
Aluminium either accumulates on the cell surface in the cell walls (Horst et al., 1999; Marienfeld et al., 2000; Wang et al., 2004) or it enters the cells (Tice et al., 1992; Lazof et al., 1994, 1996; Vázquez et al., 1999; Silva et al., 2000; Jones et al., 2006) during exposure to aluminium. However, information about the fate of aluminium associated with cell surfaces during recovery is missing. It is shown here that root cells can restore membrane functions in recovery experiments. The restoration of membrane functions together with the removal of the critical aluminium from the cell surface via its internalization and sequestering within the vacuole may contribute to the recovery of the growth. This scenario has been proposed in the present study. The vacuolar deposits in aluminium-treated maize roots support the tentative conclusion that vacuolar localization of the internalized aluminium might be the mechanism of its intracellular detoxification (Vázquez et al., 1999).
Interestingly, the high rate of aluminium internalization was typical only for meristematic cells and for the cells of the distal portion, but not of the proximal portion of the transition zone. Extracellular aluminium is mainly associated with cell wall pectins as was manifested by the correlation between the pectin content in the cell walls and the accumulation of aluminium (Horst et al., 1999; Schmohl and Horst, 2000; Hossain et al., 2006). It is speculated at this early stage that the internalization of aluminium into the cells might be closely related to the endocytosis of cell wall pectins. However, consistent with the pattern of aluminium internalization, internalization and recycling of cell wall pectins is also accomplished only in the cells of the meristem and the distal portion of the transition zone, but not in the region of rapid cell elongation (Baluka et al., 2002, 2005a; Yu et al., 2002; Paciorek et al., 2005; Dhonukshe et al., 2006). Indeed, endocytosis was active during the recovery phase as proved by the internalization of the endocytic marker FM4-64. Endocytosis proceeded in all cells of the root apex including the PTZ and the elongation zone (see FM4-64 labelling of the tonoplast), although internalization of aluminium was spatially restricted to the pectin-recycling zone (Baluka et al., 2002), and did not occur in the PTZ and the elongation zone. Inside the cells, endosomal-sorting processes might be implicated in releasing the aluminium from the pectin complexes; pectins would be recycled back to the cell wall while aluminium would continue in the endocytic pathway towards the vacuole. Pectin molecules reaching the cell wall may represent a new pool for binding of free and loosely bound apoplastic aluminium and thus support the gradual removal of morin-stained apoplastic aluminium during recovery. Supporting data for such endocytic internalization of aluminium come from the presence of aluminium in myelin figures (Vázquez, 2002), which closely resemble the multilamellar endosomes occurring in the pectin internalization pathway (Baluka et al., 2005a). Difficulties with the visualization of these intermediary structures under experimental conditions could be caused by weakening of the morin fluorescent signal intensity. A decrease of the fluorescent signal in the morin detection method after aluminium binding to pectins in vitro was shown by Eticha et al. (2005). However, this finding obtained by both the use of hand sections and fluorometric analyses of Al sorption to derived cell walls does not necessarily need to reflect the situation in intact roots of Arabidopsis exposed to morin.
Impact of aluminium on NO production and polar transport processes
The data reveal that internalization of aluminium into plant endosomes alters their behaviour as they fail to form the BFA-induced compartments. Moreover, this endosomal aluminium might also influence nitric oxide (NO) production, which showed its maximum in the cells of DTZ in control root apices but was suppressed after aluminium treatment. In animal cells, NO is active in endosomes involved in the processing of internalized heparan sulphate (Cheng et al., 2002). In addition, NO regulates endocytosis and vesicle recycling especially at neuronal synapses (Meffert et al., 1996; Huang et al., 2005; Kakegawa and Yuzaki, 2005; Wang et al., 2006). In this respect it is most interesting that plant synapses (Baluka et al., 2005b), which are very active in both endocytosis and vesicle recycling (Baluka et al., 2003, 2005b), are located exactly in the aluminium-sensitive distal portion of the transition zone (Sivaguru and Horst, 1998; Sivaguru et al., 1999).
Finally, there are obvious links between the aluminium toxicity and the inhibition of the basipetal polar transport of auxin in the epidermis and outer cortex cells. Hasenstein and Evans (1988) were the first to discover that aluminium inhibits the basipetal flow of auxin. Later, this result was fully confirmed by Kollmeier et al. (2000) who also showed that the distal portion of the transition zone is the most relevant in the aluminium-based inhibition of basipetal auxin transport. Interestingly, the cells in this zone are unique with respect to auxin and its role in cell growth regulation (Ishikawa and Evans, 1993; Baluka et al., 1994, 1996, 2004). Doncheva et al. (2005) documented that, in an aluminium-sensitive maize line, aluminium treatment mimics auxin transport inhibitors in their morphogenic effects on cell division planes in the PTZ (see also Dhonukshe et al., 2005). Polar auxin transport is insensitive to aluminium in aluminium-tolerant mutant AlRes4 of tobacco (Ahad and Nick, 2006).
The DTZ cells are not only the most sensitive towards aluminium toxicity (Sivaguru and Horst, 1998; Sivaguru et al., 1999), but are also the most active ones in the cell-to-cell transport of auxin (Mancuso et al., 2005; Santelia et al., 2005). Recently published data revealed that polar auxin transport is linked to active vesicle trafficking and that auxin is secreted out of cells via vesicle recycling (Schlicht et al., 2006). It is shown here that internalized aluminium affects the behaviour of endosomes as well as the production of NO. This indicates that the extraordinary sensitivity of the DTZ cells towards aluminium could be a consequence of an extremely active vesicle recycling driving extensive polar auxin transport in this particular root apex zone.
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Acknowledgements |
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The authors thank Milada
iamporova for critical comments on the manuscript. This work was supported in part by the Grant Agency VEGA (Grants nos 2/5085/25, 2/5086/25, and 2/3051/23). MO was supported by a Marie Curie European Reintegration Grant No. MERG-CT-2005–031168 within the 6th European Community Framework Programme. Financial support by grants from the Deutsches Zentrum für Luft- und Raumfahrt (DLR, Cologne, Germany; project 50WB 0434), from the European Space Agency (ESA-ESTEC Noordwijk, The Netherlands; MAP project AO-99–098), and from the Ente Cassa di Risparmio di Firenze (Italy) is gratefully acknowledged too. |
Abbreviations |
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BFA, brefeldin A; cPTIO, 2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide; DAF-2 DA, 4,5-diamino-fluorescein diacetate; DAG, days after germination; DMSO, dimethylsulphoxide; DTZ, distal part of the transition zone; ED, diffusion potential; Em, plasma membrane potential; EZ, elongation zone; FM4-64, (N-(3-triethylammoniumpropyl)-4-(8-(4-(diethylamino) phenyl)hexatrienyl)pyridinium dibromide); NO, nitric oxide; PTZ, proximal part of the transition zone; SHAM, salicylhydroxamic acid.
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