Download Aluminium toxicity in plants: internalization of

Journal of Experimental Botany, Vol. 57, No. 15, pp. 4201–4213, 2006
doi:10.1093/jxb/erl197 Advance Access publication 3 November, 2006
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
Peter Illéš1, Markus Schlicht2, Ján Pavlovkin1, Irene Lichtscheidl3, František Baluška1,2 and
Miroslav Ovečka1,*
1
Institute of Botany, Slovak Academy of Sciences, Dubravska cesta 14, SK-845 23, Bratislava, Slovakia
2
Institute of Cellular and Molecular Botany, University of Bonn, Bonn, Germany
3
Institution of Cell Imaging and Ultrastructure Research, University of Vienna, Vienna, Austria
Received 6 June 2006; Accepted 11 September 2006
Abstract
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 aluminiumsensitive 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 internal-
ization 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.
Introduction
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 aluminiuminduced 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; Čiamporová, 2000),
* To whom correspondence should be addressed. E-mail: [email protected]
Abbreviations: 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.
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
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4202 Illéš et al.
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 Mistrı́k, 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 (Baluška 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 (Baluška 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 noninvasive 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.
Materials and methods
Plant material and growth conditions
Seeds of Arabidopsis thaliana L., ecotype Columbia, were surfacesterilized 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 l1, calcium
pantothenate 0.1 mg l1, niacin 0.1 mg l1, pyridoxin 0.1 mg l1,
thiamin 0.1 mg l1, biotin 0.001 mg l1 of medium), FeSO4.7H2O
(1.115 mg l1), CaCl2 (111 mg l1), sucrose (10 g l1), and agar
(10 g l1), 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 lmol m2 s1 and a 12/12 h day/night
rhythm.
Aluminium sensitivity of root cells related to aluminium internalization 4203
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 lM 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 lM morin to the perfusion solution. Effects of aluminium
on Em were monitored during continual exchange of the perfusion
solution by the experimental solution (50 lM AlCl3) in perfusion
solution, perfusion speed 5 ml min1). 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 lm behind the root tip, and of the
proximal portion of the transition zone, located 300–400 lm 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 (Ovečka 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 lM AlCl3 in nutrient solution, pH 4.5, perfusion speed 10 ll
min1). After a 30 min pulse treatment, aluminium was washed out
and the roots of the Arabidopsis seedlings were stained with 100
lM 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 lM 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 lM FM4-64 was prepared from the stock solution (1 mg ml1 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 lM for 25 min. The effect of
aluminium was studied by the application of 90 lM 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 lM DAF-2 DA for
30 min and washed before observation. As a negative control, they
were treated with 10 lM 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/).
Results
Influence of aluminium on root elongation and
morphology
Arabidopsis seedlings cultivated for 7 d on agar plates with
different concentrations of AlCl3 exhibited concentrationdependent inhibition of root growth (Fig. 1). Growth of
the primary roots was only slightly reduced by 10 lM AlCl3
(to 95% of the control values, not statistically significant
according to t test at P¼0.05). Inhibition of root growth
was statistically significant at 50 lM AlCl3 as compared to
control according to t test at P¼0.01. However, growth of
the primary roots was reduced only to 89% of the control.
The inhibition was more apparent at 100 lM and 200 lM
AlCl3 (59% and 45% of control growth, respectively, highly
significant at P¼0.001), while root growth was fully inhibited
Fig. 1. Root growth of Arabidopsis plants cultivated for 7 d on agar
plates with different concentrations of AlCl3. Growth of the primary roots
is progressively affected by 100 lM and 200 lM AlCl3 while root
elongation is inhibited completely at 300 lM AlCl3. Average of 20
seedlings per treatment (6SD).
4204 Illéš et al.
Fig. 2. Effect of aluminium on elongation and morphology of the root system of Arabidopsis seedlings after 4 d cultivation. Position of root tips after
transfer of seedlings to aluminium-containing agar plates is indicated by arrows. Inhibition of root elongation at 200 lM and 300 lM AlCl3 is
accompanied by enhanced formation of lateral roots. Representative of 20 seedlings per treatment.
Fig. 3. Histochemical detection of aluminium in the roots of Arabidopsis after 7 d cultivation on agar plates by two different staining methods:
haematoxylin staining (A) and morin staining (B). Haematoxylin stained aluminium strongly at 100 lM and higher concentrations of AlCl3 while the
reaction of morin to aluminium started to be strong at 50 lM AlCl3. Note the radial expansion of root cells at 100 lM and 200 lM AlCl3, but not at
300 lM AlCl3 due to the complete inhibition of root growth. Representative of five seedlings per treatment. Bar¼100 lm.
by 300 lM AlCl3 (only 2% root elongation as compared
to control plants, highly significant at P¼0.001; Figs 1, 2).
Severe inhibition of root elongation was accompanied by radial expansion of the root cells at 100 lM and 200 lM AlCl3,
but not at 300 lM AlCl3, due to the complete inhibition of root
growth (Fig. 3). Root architecture of Arabidopsis seedlings
exposed to lower concentrations of aluminium (10–100
lM AlCl3) was almost unaffected. However, 200 lM and
Aluminium sensitivity of root cells related to aluminium internalization 4205
300 lM 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.
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 lM AlCl3 (data
not shown), aluminium staining by haematoxylin was
not detectable and only a weak diffuse fluorescence of morin
occurred. At 50 lM 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 lM and 200 lM
AlCl3, both staining methods showed maximum accumulation of aluminium in the root apex. In the plants exposed to 300 lM 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 aluminiumsupplemented agar plates. The critical discrimination limit
in the sensitivity between morin and haematoxylin is apparently at 50 lM AlCl3 (Fig. 3). Because of mild effects on
root growth and the capability of morin to localize aluminium, 50 lM 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 lm behind the root tip, the proximal part of the
transition zone 300–400 lm behind the root tip (Verbelen
et al., 2006). Em of cortical cells varied between 82 mV
and 98 mV (8864 mV, n¼37) in the distal part of the
transition zone (DTZ) and between 94 mV and 117 mV
(10567 mV, n¼29) 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. energydependent, 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 lM 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.
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 lM 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
Fig. 4. Effect of 1 mM NaCN and 1 mM salicylhydroxamic acid
(SHAM) on cortical cell membrane potential (Em). Both in proximal
transition zone (A) and distal transition zone (B), the Em rapidly
depolarized to the values of diffusion potential (ED). Representative of
20 seedlings per treatment.
4206 Illéš et al.
Fig. 5. Changes of cortical cell membrane potential (Em) after treatment
with 50 lM AlCl3. Em depolarized rapidly (within 2 min) after aluminium
application; depolarization was more extensive in the cells of distal
transition zone (DTZ) than in the proximal transition zone (PTZ). After
removing aluminium, Em in the PTZ completely repolarized within 3 min,
while in the DTZ within 10 min (A). After 30 min aluminium treatment,
the complete repolarization of Em occurred within 6 min in the cells of
PTZ and within 14 min in the DTZ (B). Representative of 24 seedlings
per treatment.
was fully reversible, which was consequently followed by
recovery of root growth (data not shown).
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 lM 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 cyto-
plasm 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).
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; Ovečka et al.,
2005; Šamaj 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).
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;
Šamaj 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.
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
Aluminium sensitivity of root cells related to aluminium internalization 4207
Fig. 6. Time-course of aluminium uptake in the cells of the meristem and DTZ monitored by morin. Pulse treatment of Arabidopsis roots with 50 lM
AlCl3 for 30 min, followed by washing and morin staining. Observation of the root cells 20 min after the end of aluminium treatment revealed the
presence of aluminium only in the apoplast (A). The first signals of morin fluorescence in the cytoplasm were detected within 1 h (B). 2 h and 30 min after
treatment aluminium accumulated into roundish vacuole-like structures of varying size (C). 3 h 30 min after treatment aluminium was sequestered in
vacuole-like compartments (D). Representative of five seedlings per treatment. Bar¼10 lm.
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 lM 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).
Discussion
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; FischerParton et al., 2000; Bolte et al., 2004; Ovečka et al., 2005;
Šamaj 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
4208 Illéš et al.
Fig. 7. Internalization of aluminium in specific developmental zones of pulse-treated Arabidopsis roots with 50 lM AlCl3 for 30 min. Roots were
pretreated with 4 lM FM4-64 and stained with morin after washing out the aluminium. In the meristematic cells (A) and the cells of distal transition zone
(B) aluminium was internalized and accumulated in vacuolar compartments after 2 h 50 min of recovery. Tonoplast was labelled red with FM4-64 and
the aluminium-containing lumen of the vacuole was stained green with morin. In the proximal transition zone (C) there was no uptake of aluminium even
3 h 10 min after the end of the treatment; aluminium was not accumulated in the vacuoles and could be detected only in the apoplast. Representative of
five seedlings per treatment. Bar¼10 lm.
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 (Baluška
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
Aluminium sensitivity of root cells related to aluminium internalization 4209
Fig. 8. Effect of aluminium on internalization of endocytic marker FM4-64. Control root after 10 min labelling with FM4-64 dye (A). Formation of
BFA-induced compartments by 35 lM BFA in FM4-64-labelled roots (B). Treatment with 90 lM aluminium for 90 min did not change considerably the
pattern of FM4-64 labelling (C), but it prevented formation of BFA-induced compartments after application of 35 lM BFA (D). Representative of five
seedlings per treatment. Bar¼10 lm.
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 sup-
port 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 (Baluška 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
4210 Illéš et al.
Fig. 9. Detection of NO production by DAF-2DA labelling in control root tip (A), root tips treated with 90 lM aluminium for 60 min (B), and 10 lM
cPTIO, the NO-scavenger, for 60 min (C). Fluorescence of DAF-2DA is green, FM4-64 is red. Fluorescence intensity (D) and distribution (insert) of
DAF-2DA labelling along root developmental zones. Note the disappearance of the NO production peak in DTZ after aluminium treatment. Average
intensities of 20 roots per treatment.
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 pectinrecycling zone (Baluška 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 (Baluška
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
Aluminium sensitivity of root cells related to aluminium internalization 4211
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 (Baluška et al., 2005b), which
are very active in both endocytosis and vesicle recycling
(Baluška 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; Baluška 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.
Acknowledgements
The authors thank Milada Čiamporova 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 AO99–098), and from the Ente Cassa di Risparmio di Firenze (Italy) is
gratefully acknowledged too.
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