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Plant Physiol. (1998) 118: 159-172
Alterations in the Cytoskeleton Accompany Aluminum-Induced Growth
Inhibition and Morphological Changes in Primary Roots of
Maize1
Elison B. Blancaflor2,
David L. Jones2, and
Simon Gilroy*
Biology Department, 208 Mueller Laboratory, The Pennsylvania State
University, University Park, Pennsylvania 16802 (E.B.B., S.G.); and School of Agricultural and Forest Sciences, University of Wales,
Bangor, Gwynedd, United Kingdom (D.L.J.)
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ABSTRACT |
Although
Al is one of the major factors limiting crop production, the mechanisms
of toxicity remain unknown. The growth inhibition and swelling of roots
associated with Al exposure suggest that the cytoskeleton may be a
target of Al toxicity. Using indirect immunofluorescence microscopy,
microtubules and microfilaments in maize (Zea mays
L.) roots were visualized and changes in their organization and stability correlated with the symptoms of Al toxicity.
Growth studies showed that the site of Al toxicity was associated with
the elongation zone. Within this region, Al resulted in a
reorganization of microtubules in the inner cortex. However, the
orientation of microtubules in the outer cortex and epidermis remained
unchanged even after chronic symptoms of toxicity were manifest.
Auxin-induced reorientation and cold-induced depolymerization of
microtubules in the outer cortex were blocked by Al pretreatment. These
results suggest that Al increased the stability of microtubules in
these cells. The stabilizing effect of Al in the outer cortex coincided
with growth inhibition. Reoriented microfilaments were also observed in
Al-treated roots, and Al pretreatment minimized cytochalasin B-induced
microfilament fragmentation. These data show that reorganization and
stabilization of the cytoskeleton are closely associated with Al
toxicity in maize roots.
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INTRODUCTION |
The trivalent metal Al is solubilized by the low pH of acidic
soils and is a major factor limiting crop productivity in many areas of
the world. One of the most obvious symptoms of Al toxicity is the rapid
inhibition of root growth (Ryan et al., 1993 ), which results in poor
nutrient acquisition and consequently leads to nutrient deficiencies
and decreased crop yields (Taylor, 1990; Kochian, 1995). Despite many
proposed theories attempting to explain the cause of toxicity, however,
the mechanisms involved remain unidentified (Kochian, 1995; Kochian and
Jones, 1997).
The cytoskeleton is a dynamic network of proteinaceous components
consisting mainly of microtubules and microfilaments. Both of these
cytoskeletal components have been implicated in a wide variety of
cellular functions, such as cell division, cell expansion, cell wall
synthesis, organelle movement, and tip growth (Seagull, 1989).
Compounds that disrupt the normal functioning of the cytoskeleton have
been shown to inhibit growth and cause substantial swelling of the root
apex, similar to the visual symptoms of Al rhizotoxicity (Baskin et
al., 1994; Baluska et al., 1996; Baskin and Wilson, 1997). In addition,
several factors that lead to altered growth in roots, such as
gravistimulation, osmotic stress (Blancaflor and Hasenstein, 1993,
1995a), and hormone treatments (Baluska et al., 1993a; Blancaflor and
Hasenstein 1995b), elicit a reorientation in the microtubule
cytoskeleton. Therefore, the growth inhibition and concomitant increase
in root diameter observed in roots exposed to Al (Bennet et al., 1985;
Ryan et al., 1993; Sasaki et al., 1996) suggest that the plant
cytoskeleton could be a cellular target of Al phytotoxicity.
Various neurological disorders induced by Al in experimental animals
have also been associated with abnormalities in components of the
cytoskeleton (Schmidt et al., 1991; Singer et al., 1997). One proposed
model for such cytoskeletal disruption involves Al binding
107-fold more effectively than Mg to the
GTP-binding sites on tubulin that are required for microtubule
assembly. When Al is associated with the GTP-tubulin complex, the
exchange of GTP for GDP is slower. Thus, the intricate dynamics of
microtubule formation could be disrupted (MacDonald et al., 1987;
MacDonald and Martin, 1988). In addition, the study of Al-induced
neurofibrillary tangles in mammalian systems has shown that Al can also
affect the expression of cytoskeletal regulatory genes, cytoskeletal
protein phosphorylation, and the production of secondary messengers
(cAMP, cGMP, inositol 1,4,5-trisphosphate) involved in the regulation
of cytoskeletal dynamics (Strong et al., 1997).
Control of plant cytoskeletal organization has been suggested to be
under the influence of cellular events and mediated by one or more
signal transduction pathways (Cyr and Palevitz, 1995). The
phosphoinositide signaling system (Xu et al., 1992; Drobak, 1993) and
Ca2+/calmodulin (Cyr, 1991; Fisher et al., 1996)
have been implicated in the regulation of the plant cytoskeleton and
have also been proposed as cellular targets of Al toxicity (Siegel and
Haug, 1983; Jones and Kochian, 1995, 1997; Jones et al., 1998).
Despite the possibility that the cytoskeleton could be a principal
target of Al rhizotoxicity, there have been only a few studies
investigating the effects of Al on the plant cytoskeleton. Although it
has been shown that Al strongly promotes the polymerization of tubulin
subunits into microtubules in vitro (MacDonald et al., 1987; MacDonald
and Martin, 1988), microtubules in elongating cells of wheat roots were
reported to depolymerize in response to Al (Sasaki et al., 1997).
Grabski and Schindler (1995) reported that Al induced a rigidification
of the actin network in soybean root suspension cells, and it was
recently shown that mRNAs of the actin-bundling protein fimbrin were
up-regulated in wheat roots exposed to Al (Ortega et al., 1997). Al has
also been shown to inhibit growth in liverwort rhizoids concomitant
with a fragmentation of the actin network (Alfano et al., 1993).
The studies cited above clearly indicate that Al could potentially
affect the plant cytoskeleton; however, these experiments do not
provide a consistent account of this effect. Thus, it is still
uncertain whether Al acts by stabilizing, depolymerizing, or causing a
reorganization in the cytoskeleton. Furthermore, the link between
changes in the cytoskeleton and Al toxicity symptoms in intact roots
remains unproved. Therefore, we investigated the effects of Al on the
microtubules and microfilaments in elongating cells of maize roots and
correlated them with Al-induced growth inhibition. We present data
showing that microtubules and microfilaments are altered in both
stability and organization when exposed to Al. The data show that Al
interactions with cytoskeletal components may be an important factor in
Al rhizotoxicity.
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MATERIALS AND METHODS |
Plant Material
Seeds of maize (Zea mays L. cv Merit, Asgrow Seed Co.,
Kalamazoo, MI) were soaked in distilled water for 10 h and
germinated between wet paper towels. Seedlings were grown vertically
for 2 d under fluorescent lamps providing continuous diffuse light (36 µmol m 2 s1).
Seedlings with straight roots approximately 2 cm in length were
selected and transferred to 350-mL clear, plastic chambers containing
aerated 200 µM CaCl2, pH 4.5. Roots
were grown hydroponically at 25°C in this solution for 24 h
before experimentation.
Growth Measurements and Al Treatments
Straight roots, approximately 3 to 4 cm in length, were selected
and marked with black oil-based Speedball ink at 1-mm intervals from
the root tip. One-half of the marked roots were returned to the
solution containing 200 µM CaCl2,
pH 4.5 (controls), and the other half were placed in a solution
containing 200 µM CaCl2, 50 µM AlCl3, pH 4.5 (Al3+ activity = 27.8 µM
assuming no Al(OH)3 precipitation). The length of
each segment was measured from digitized images of the marked roots
2 h after treatment. In another set of growth measurements, elongation of the whole root was measured for 12 h at 20-min
intervals. For root-diameter changes, images of roots were captured
every 2 h for 10 h. Root diameter was measured at 1-mm
intervals along the length of the root, with the tip of the cap
designated as 0. Images of roots were collected at each time point
using a 100-mm Promaster Macro lens (Nikon) attached to a video camera
(model C2400, Hamamatsu, Tokyo, Japan) and captured with a LG-3 frame grabber (Scion Corp., Frederick, MD) and Quadra 800 computer (Apple Computer Inc., Cupertino, CA) running IPLabs Spectrum image-acquisition software (Signal Analytics, Vienna, VA).
Light Microscopy of Roots
Plants were grown hydroponically as described above and treated
with or without Al (50 µM) for 24 h. The terminal 7 mm of the roots was then excised and fixed under a partial vacuum in 3% (w/v) glutaraldehyde, 50 mM sodium phosphate buffer, pH
7.2, for 24 h. After fixation, tissue samples were dehydrated
through an ethanol series (25%, 50%, 75%, and 100% [v/v] ethanol
for 12 h each), exchanged with acetone (100%, v/v), exchanged
back to ethanol (100%, v/v), and resin embedded using a historesin
embedding kit (Leica Instruments, Heidelberg, Germany). Embedded roots
were then sectioned (4 µm thick) on a microtome (model 2050, Reichert-Jung, Heidelberg, Germany) using glass and tungsten-carbide
steel knives, stained for 2 h in toluidine blue (0.05% [w/v] in
benzoate buffer, pH 4.0), and visualized using a LSM410 confocal
microscope (Zeiss).
Assessment of Cell Viability
To confirm the viability of root cells, plants were exposed to Al
(50 µM) for 24 h as described above and then stained
with FDA (0.05% [w/v]; Sigma) for 5 min as described by Huang et al. (1986). After washing (1 min), the fluorescence of cells within the
elongation zone of control and Al-treated roots was then imaged with a
confocal microscope, with excitation at 488 nm and emission at 515 to
540 nm.
Immunofluorescence
Indirect immunofluorescent labeling of microtubules and
microfilaments was essentially performed as described by Blancaflor and
Hasenstein (1993, 1997) with minor modifications. After the experimental treatments, the terminal 7 mm of the roots was excised and
placed into glass vials containing fixative. For preserving microtubules the fixative consisted of PHEMD buffer (60 mM
Pipes, 25 mM Hepes, 10 mM EGTA, 2 mM MgCl2, and 5% [v/v] DMSO), pH
7.0, plus 4% (w/v) formaldehyde (Blancaflor and Hasenstein, 1993), whereas fixative for preserving microfilaments consisted of 2% formaldehyde in PHEMD buffer, pH 7.0 (Blancaflor and Hasenstein 1997).
After 2 h in fixative, roots were washed (3 × 5 min
each) in PHEMD buffer, attached to a metal block with cyanoacrylate (Krazy Glue, Borden Inc., Columbus, OH), and sectioned
longitudinally with a Vibratome-1000 (Technical Products International,
St. Louis, MO) at a thickness of 80 µm. Median longitudinal sections
were transferred to glass slides coated with poly-L-Lys
(Sigma) and allowed to dry for 5 min. Sections were partially digested
for 10 min in an enzyme solution consisting of 1% (w/v) cellulase (Yakult, Tokyo, Japan), 0.01% (w/v) pectolyase (Sigma), and 1% (w/v)
BSA followed by 15 min of incubation in 1% (v/v) Triton X-100.
Sections were then incubated in the primary antibodies for 2 h.
For labeling microtubules, sections were incubated in rat anti-yeast
tubulin (clone YOL 1/34, Accurate Chemical and Scientific Corp.,
Westbury, NY). For labeling actin, a mouse monoclonal antibody to
phalloidin-stabilized pea actin was used (Andersland et al., 1994).
Sections were then incubated in the dark with the appropriate secondary
antibodies conjugated to fluorescein isothiocyanate (goat anti-rat IgG
for microtubules and rabbit anti-mouse IgG for microfilaments; Sigma).
After three washes in PHEMD buffer, sections were mounted in 20%
Mowiol 4-88 (Calbiochem, La Jolla, CA) in PBS, pH 8.5, containing
0.1% pphenylenediamine (Sigma). Cytoskeletal elements
were imaged with a confocal microscope, with excitation at 488 nm and
emission at 515 to 540 nm. Images from the confocal microscope were
assembled using Photoshop 3.0 (Adobe Systems, Inc., Mountain View, CA)
and printed on a dye-sublimation printer (Phaser II SDX, Tektronix,
Inc., Wilsonville, OR).
Auxin, Cold, Taxol, and CB Treatment
A stock solution of 10 mM IAA (Sigma) was prepared in
2-propanol. CB (Calbiochem) was prepared as 10 mM stock
solutions in 100% DMSO. The required volumes of IAA and CB stock were
added to separate sets of roots growing in 200 µM
CaCl2, pH 4.5, with or without 50 µM Al. The final concentration of IAA in the solution was
1 µM, and for CB, 50 µM. IAA-treated roots
were fixed and processed for microscopy after 1 h of incubation,
whereas CB-treated roots were processed after 3 h. For cold
treatment, a solution containing 50 µM Al was prechilled
to 2°C. Roots grown at 25°C for up to 12 h in solutions
containing Al were transferred to 2°C solutions for 2 h and
processed for microscopy.
For taxol experiments, a stock of 20 mM taxol (Fluka) was
prepared in 100% DMSO. A working solution of 20 µM taxol
was made by adding the required volume of the stock solution to 200 µM CaCl2, pH 4.5. Roots were fixed
after 12 and 48 h of incubation in taxol and processed for
microscopy. In another set of experiments roots were incubated in taxol
solution for 3 h, transferred to a solution containing 1 µM IAA for 1 h, and processed for microscopy. Solvent controls (0.5% [v/v] DMSO or 0.01% [v/v] propanol) showed no effect on growth or cytoskeletal organization.
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RESULTS |
Effects of Al on Growth, Morphology, and Viability of Maize Root
Cells
Growth rates of maize roots were inhibited by Al in a
concentration-dependent manner. Less than 20% inhibition occurred at 5 µM, and greater than 60% inhibition occurred at
concentrations at or above 50 µM (Fig.
1A). Because maximum root-growth
inhibition under our conditions occurred at Al concentrations of 50 µM or greater (Fig. 1A), all subsequent experiments were
performed on roots grown hydroponically in the presence or absence of
50 µM Al. The growth of maize primary roots proceeded at
a rate of approximately 1.5 mm h1 under control
conditions. Application of 50 µM Al resulted in a rapid
decline in the growth rate, consistent with results presented previously for Al-sensitive varieties of wheat (Jones and Kochian, 1995) and maize (Sivaguru and Horst, 1998). The root-growth rate was
reduced by 50% after 2 h and by 80% after 12 h (Fig. 1B), with a complete inhibition of growth evident after 24 h of Al exposure (data not shown).
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| Figure 1.
Effects of Al on elongation and radial expansion
of maize roots. A, Dose-response curve of root-growth inhibition by Al.
Seedlings were transferred to aerated solutions containing the
indicated concentrations of Al and the root growth was measured after
6 h. Concentrations at or above 50 µM Al resulted in
maximum growth inhibition. B, Kinetics of axial root-growth rate of
untreated roots (control) and roots in response to 50 µM
Al. The growth rate of roots declined within 1 h of Al application
(arrow). C, Effect of 50 µM Al on elongation growth
within different regions of the root. Roots were marked at 1-mm
intervals from the tip, and extension of the marked segments was
measured 2 h after exposure to Al. Growth of the region 2 to 5 mm
from the root tip was the most effectively inhibited by Al. D, Kinetics
of radial expansion at 2 and 4 mm from the tip of maize primary roots
treated with 50 µM Al. Swelling was perceptible more than
3 mm from the tip 6 h after Al exposure. No swelling was observed
less than 2 mm from the tip during the initial 10 h of Al
treatment. Data points are means from six or more roots ± SE.
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Growth inhibition after Al exposure was most pronounced in the region 2 to 5 mm from the root tip after 2 h (Fig. 1C). This pattern was
maintained for as long as we monitored the roots (6 h; data not shown)
and was similar to the inhibitory effect of Al reported for squash
roots (Le Van et al., 1994). For this reason, all later studies
concentrated on the zone of elongation 2 to 5 mm from the root cap. In
addition to the rapid inhibition of root growth, Al also induced an
increase in root diameter over longer periods (>4 h); however, this
appeared to be limited to distances from 3 to 5 mm from the root cap
(Fig. 1D). Prolonged exposure to Al (>12 h) also resulted in severe
morphological distortions in the root-elongation zone, as shown by
swollen cells within the inner cortex and large holes in the root
surface produced by the loss of both epidermal and outer cortical
groups of cells (Fig. 2, A and B; see
also Fig. 6). Despite an obvious degeneration of the root cortex,
studies with the vital stain FDA showed no loss of cell viability in
control or Al-treated cells in the cortical and epidermal regions,
indicating that Al does not act as a generic cytotoxin (Fig. 2, C and
D).
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| Figure 2.
Light micrographs and viability staining of maize
primary roots after exposure to 50 µM Al. Roots were
fixed and embedded in historesin, and a median longitudinal section was
taken from control roots (A) and roots 24 h after Al treatment
(B). Inset in A shows a magnified view of the surface of an untreated
root. Inset in B shows a magnified view of the surface lesions of an
Al-treated root. These lesions were not generated by a sectioning
artifact, as shown in Figure 6. Confocal sections of the fluorescence
from cells at the surface of control (C) and Al-treated (D) roots
stained with the viability stain FDA show that despite the degeneration
of the root cortex and holes (arrow) in the surface layer of cells, the
cells at the surface of the Al-treated root are viable. Images are
representative of at least 50 independent root samples. Bar in B = 1 mm; bars in insets = 100 µm; and bar in D = 25 µm and
applies to C and D.
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| Figure 6.
Morphology of maize primary roots after exposure
to 50 µM Al or 20 µM taxol. Like Al-treated
roots, taxol-treated roots showed disintegration of outer cell layers
(arrows) compared with untreated roots (control). By 24 h swelling
was no longer apparent because portions of the epidermis and outer
cortex had sloughed off. Bar = 1 mm.
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Effect of Al on Microtubule Organization
The organization of microtubules in primary roots of maize exposed
to 50 µM Al for various periods (1-24 h) was examined by indirect immunofluorescence microscopy. In control roots cortical microtubules in the elongation zone were oriented perpendicular to the
long axis of the root in the outer cortex (Fig.
3A), inner cortex (Fig. 3B), and stele
(Fig. 3C). This pattern of microtubule orientation was observed
throughout the apical 6 mm of the root. At 6 to 7 mm from the root tip,
cortical microtubules shifted to an oblique orientation typical of all
cell layers in the maturation zone (Fig. 3D). These results are
consistent with those of previous studies (Blancaflor and Hasenstein,
1993, 1995b).
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| Figure 3.
Organization of cortical microtubules in maize
primary roots grown in 200 µM CaCl2, pH 4.5 (controls). Along the elongation zone (approximately 2-5 mm from the
root tip), the outer (A) and inner (B) cortex of control roots show
microtubules aligned perpendicular to the long axis of the root. C,
Stelar cells in the same region also show transverse microtubules.
D, Microtubules shift to an oblique orientation 6 to 7 mm from
the root tip. The schematic diagram of the root indicates the regions
where immunofluorescence images were obtained. Symbols in parentheses
in A to D correspond to the symbols in the root diagram. Images are
representative of at least five independently processed root samples.
Bar in A = 25 µm.
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Upon initial exposure to Al (1-2 h), the organization of microtubules
in the elongation zone (2-5 mm from the root cap) did not differ from
that in control roots (data not shown); however, after 3 h of Al
exposure, cells in the three layers of the inner cortex 4 to 5 mm from
the root cap had started to display microtubules oriented in either a
random or an oblique pattern (Fig. 4A).
This zone of reorientation moved progressively closer to the root apex upon prolonged exposure to Al, with microtubules in the inner cortex of
the entire elongation zone becoming reoriented by 12 h (Fig. 4B).
After 4 to 5 h of Al exposure, the pattern of microtubules in the
stele had also changed from their typically ordered and transverse
orientation to being primarily longitudinal (compare Figs. 3C and 4C).
Despite the reorganization of microtubules in the inner cortex and
stele, the orientation of microtubules in cells of the outer cortex and
epidermis remained principally transversely oriented after 4 h
(Fig. 4D) and even after prolonged exposure (>12 h; Fig. 4E). Although
microtubules of inner cortical cells in the elongation zone were
clearly reorganized after Al treatment, microtubules of all cell layers
in the maturation zone did not appear to be affected by Al and remained
similar to those of control roots (compare Figs. 3D and 4F).
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| Figure 4.
Organization of cortical microtubules in maize
primary roots after exposure to 50 µM Al. A, After 3 h of continuous exposure to Al, cells in the inner cortex 4 to 4.5 mm
from the root tip showed random and obliquely oriented microtubules. B,
After 12 h of continuous exposure to Al, reoriented microtubules
occurred closer to the root tip. Cells in the inner cortex 2 mm from
the tip showed random to longitudinally oriented microtubules. Arrow
shows region where outer cortex has sloughed off. C, The stelar cells
also exhibited random to longitudinal microtubules but occurred after
4 h of Al exposure. D, Four hours after Al exposure, outer
cortical cells 4 mm from the root tip retained their net transverse
microtubule orientation. E, Outer cortical cells 12 h after Al
exposure also retained an overall transverse alignment of microtubules
despite the distorted appearance of the cells. F, Cells in the
maturation zone (about 6 mm from the tip) showed oblique microtubules
that were similar to controls. The schematic diagram of the root
indicates the regions where immunofluorescence images were obtained.
Symbols in parentheses in A to F correspond to the symbols in the root
diagram. Images are representative of at least five roots per time
point. Bar in F = 25 µm.
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Al Prevents Auxin-Induced Microtubule Reorientation and
Cold-Induced Microtubule Depolymerization
In addition to affecting the organization of microtubules within
cells, Al may also be able to affect the stability of cytoskeletal elements either directly or indirectly. This is supported by in vitro
studies in which Al has been shown to promote tubulin assembly into
microtubules (MacDonald et al., 1987). To determine whether the
stability of microtubules is affected by Al, we exposed roots to agents
known to reorient or depolymerize microtubules either before or after
pretreatment with Al.
Auxin has been shown to cause microtubules in cortical cells of maize
roots to reorient from a primarily transverse to a longitudinal pattern
(Blancaflor and Hasenstein, 1995b), and has also been reported to
induce the depolymerization of microtubules (Baluska et al., 1996). To
test the possibility that Al may be altering either the stability or
the ordering of microtubules, roots were incubated with or without Al
before treatment with auxin.
Roots grown in the absence of Al followed by a 1-h exposure to 1 µM IAA displayed a characteristic reorientation of
microtubules from transverse to longitudinal arrays in both the outer
(Fig. 5A) and inner cortex (data not
shown) of the elongation zone, in agreement with Blancaflor and
Hasenstein (1995b). In contrast, roots incubated in 50 µM
Al for 1 h before IAA application did not show an IAA-induced
reorientation of the microtubules in the outer cortex (Fig. 5B), but
did show the reorientation of microtubules in the inner cortex to
principally longitudinal arrays (data not shown).
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| Figure 5.
Organization of cortical microtubules in maize
primary roots treated with IAA, Al, or cold. A, Microtubules in cells
of the outer cortex shifted to longitudinal orientations after 1 h
of exposure to 1 µM IAA. B, Microtubules in cells of the
outer cortex exposed for 1 h to 50 µM Al followed by
1 h in 1 µM IAA remained transversely oriented. C,
Roots incubated in 2°C solution for 2 h showed fragmented
microtubules. D, Roots pretreated with Al for 12 h before cold
exposure still had cells with intact microtubules. E, The fraction of
cells in the outer cortex of cold-treated and Al-plus-cold-treated
roots were classified by whether they had intact or
fragmented/depolymerized microtubules. At least 50 cells were
observed for each treatment. Images are representative of at least
three roots from three independent experiments. The schematic diagram
of the root indicates the regions where immunofluorescence images were
obtained. Symbols in parentheses in A to D correspond to the symbols in
the root diagram. Bar in B = 25 µm.
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Another treatment known to depolymerize microtubules in roots is cold
temperatures (Baluska et al., 1993b). Microtubules of roots incubated
at 2°C for 2 h were extensively fragmented (Fig. 5C). Roots
pretreated for 3 h with Al before cold application had a greater
density of intact microtubules (Fig. 5D). This effect was
representative of up to 12 h of Al pretreatment. Only 39% of
cells in the outer cortex of roots exposed to cold had intact microtubules. However, in roots pretreated with Al before cold application, 87% of outer cortical cells had intact microtubules (Fig.
5E).
To further test the interaction between Al and cytoskeletal elements,
we compared the organization of microtubule arrays after treatment with
taxol, a drug known to promote microtubule assembly and stabilize
preexisting microtubules in plant cells (Bokros et al., 1993; Chu et
al., 1993), with those of Al-treated roots. Our rationale for doing
this was based on observations showing that the morphology of roots
exposed to Al was visually similar to that of roots treated with taxol.
Taxol-treated roots exhibited a distinct swelling along the elongation
zone and eventually developed lesions along the surface of the root,
similar to those seen under Al toxicity. However, these symptoms took
longer (i.e. >48 h) to develop than was observed for Al (24 h; Fig.
6).
Examination of microtubules in the outer cortex of 12-h taxol-treated
roots revealed bundled, dense, and highly ordered microtubule arrays
(Fig. 7A). Although 12 h of Al
exposure did not result in increased ordering of microtubules, bundling
was more extensive compared with controls, as shown by the increased
lateral association between microtubules (Fig. 7B). Control roots
exhibited bundled microtubules, but the degree of bundling was lower
(Fig. 7C; see also Fig. 1). After 48 h, the cells in the cortex of
taxol-treated roots were no longer elongated but displayed distorted
shapes similar to those of Al-treated roots. Despite the distorted
appearance of these cells, an overall transverse alignment of
microtubules was still apparent in both the outer (Fig. 7D) and inner
cortex (data not shown). Microtubules in outer cortical cells of 24-h Al-treated roots also retained an overall transverse microtubule alignment with extensive bundling (Fig. 7E). Roots treated with 20 µM taxol for 3 h followed by 1 h in 1 µM IAA exhibited transverse microtubules in the outer
cortex (Fig. 7F).
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| Figure 7.
Comparison of cortical microtubule organization in
maize primary roots treated with taxol or Al. A, Microtubules in taxol
(20 µM)-treated roots were higher in density and
displayed extensive bundling compared with controls. Bundling was
characterized by an increased lateral association of microtubules. B,
Although not as dense as microtubules in taxol-treated roots, 50 µM Al caused the formation of bundled microtubules
(arrowheads). For comparison, the low degree of bundling in untreated
controls is shown in C. D, Outer cortical cells after 48 to 60 h
of exposure to 20 µM taxol showed an overall transverse
alignment of microtubules despite distorted cell shapes. E, Region
where the outer cortical cells in the elongation zone remained intact
even after 24 h of exposure to 50 µM Al. Like
taxol-treated roots, microtubules retained an overall transverse
alignment and displayed extensive bundling (arrowhead). F, Cortical
microtubules of roots incubated in taxol for 3 h followed by
1 h of incubation in 1 µM IAA remained dense and
transversely oriented. Images are representative of at least five roots
per treatment. The schematic diagram of the root indicates the regions
where immunofluorescence images were obtained. Symbols in parentheses
in A to F correspond to the symbols in the root diagram. Bar in E = 10 µm and applies to A to E; bar in F = 25 µm.
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Effects of Al on Microfilament Organization
Cells in the elongation zone of untreated control roots showed
microfilaments predominantly longitudinal in orientation present in
both the outer and inner cortex, as well as in the stele parenchyma (Fig. 8, A-C). There were no gross
qualitative changes in the organization of microfilaments treated with
Al during the initial 5 h of exposure (data not shown). However,
greater than 6 h of Al exposure resulted in microfilaments in the
inner cortex becoming more random in orientation (Fig. 8D). In
contrast, microfilaments in the outer cortical cells remained
longitudinal in orientation and similar to controls (Fig. 8E).
Disruption of the actin network in the inner cortex was severe after 8 to 24 h of Al treatment. Thick actin cables were randomly oriented
and appeared to radiate from the nucleus (Fig. 8F); however, finer
strands of randomly oriented actin bundles were also present throughout
the cell (Fig. 8G). Outer cortical cells after 24 h of exposure to
Al also showed thicker microfilament cables but the predominant
alignment was still in the longitudinal direction, as in nontreated
cells (Fig. 8H). Despite a disruption of microfilament organization in
cells of the inner cortex within 6 to 24 h of Al exposure,
microfilaments in the stelar parenchyma cells remained longitudinal.
However, the microfilament bundles in these cells appeared thicker than those in controls (Fig. 8I).
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| Figure 8.
Organization of actin microfilaments in maize
roots exposed to 50 µM Al. Microfilaments in the inner
cortex (A), outer cortex (B) and stele (C) of control roots are
parallel to the long axis of the cell. D, By 6 h of Al exposure,
the inner cortical cells show a more random organization of
microfilament bundles. Compared with the controls, microfilament
bundles are not as straight (arrows). E, After 6 h the outer
cortical cells still show a preferential longitudinal alignment of
microfilaments. F, Microfilaments in the inner cortex after 12 to
24 h of Al exposure are randomly oriented. Thick microfilament
bundles (arrow) appear to radiate from the nucleus (n).
G, Higher-magnification image of an inner cortical cell after 24 h
of Al exposure showing randomly oriented microfilaments. H, Outer
cortical cells 24 h after Al treatment still show an overall
longitudinal orientation of microfilaments, but microfilament bundles
were generally thicker. I, Despite the disruption of microfilament
organization in the other tissues, microfilaments in the stele were
still longitudinal in orientation after a 24-h exposure to Al. The
schematic diagram of the root indicates the regions where
immunofluorescence images were obtained. Symbols in parentheses in A to
I correspond to the symbols in the root diagram. Bar in C = 25 µm and applies to A to E and I; bar in F = 25 µm and applies
to F and H; and bar in G = 10 µm.
|
|
To test whether Al can induce a stabilization of microfilaments
in intact maize tissues, roots were pretreated with Al before application of the known actin-depolymerizing compound CB. Exposure of
roots to 50 µM CB for 3 h resulted in an extensive
fragmentation of the actin network in cortical cells throughout the
elongation zone (Fig. 9A), with some
lesser fragmentation evident in the stele. Remaining microfilament
bundles were also thinner and less dense in the stele of CB-treated
roots compared with controls (Fig. 9B). These results are consistent
with those of a previous study (Blancaflor and Hasenstein, 1997). Roots
pretreated with Al for 3 to 12 h before CB application also
exhibited some limited fragmentation of the actin network; however,
many cells in both the inner and outer cortex retained a high density
of thick microfilament cables (Fig. 9C). Microfilaments in the stele
were also more resistant to the effects of CB. Microfilament bundles
remained dense and thicker than in roots not treated with Al (Fig. 9D).
In roots treated with CB only 15% of cells in the outer cortex had an
intact array of microfilaments. Incubation in Al for 3 h before CB
application resulted in a higher percentage (68%) of cells in the
outer cortex with thick bundles of microfilaments (Fig. 9E).
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| Figure 9.
Organization of actin microfilaments in maize
roots treated with the actin-depolymerizing agent CB (A and B) or Al
plus CB (C and D). A, Microfilaments in the outer cortical cells of
maize roots exposed to 50 µM CB for 3 h were
completely disrupted. Only a few thick microfilament bundles remained
(arrow). B, Microfilaments in the stele were also disrupted after CB
treatment and characterized by thinner bundles. C, Cells in the outer
cortex of roots pretreated with Al for 3 h still retained numerous
thick microfilament bundles (arrows) even after exposure to CB. D,
Microfilament bundles in the stele of Al-pretreated roots were thick
and dense despite 3 h of exposure to CB. E, The fraction of cells
in the outer cortex of CB-treated and Al-plus-CB-treated roots were
classified by whether they had intact microfilaments or fragmented/no
microfilaments. At least 50 cells were observed for each treatment.
Images are representative of at least three roots from three
independent experiments. The schematic diagram of the root indicates
the regions where immunofluorescence images were obtained. Symbols in
parentheses in A to D correspond to the symbols in the root
diagram. Bar in A = 25 µm.
|
|
| |
DISCUSSION |
Despite the advancement of many theories to explain the cause of
Al toxicity in plants, the initial target sites still remain unknown
(Kochian, 1995; Kochian and Jones, 1997). It has recently been
hypothesized from experiments with cell-suspension cultures that one of
the primary mechanisms of Al toxicity in plants may be associated with
an increased rigidity of the actin network (Grabski and Schindler,
1995). In addition, Al has also been shown to promote the assembly of
microtubules in vitro (MacDonald et al., 1987). However, there are also
conflicting reports suggesting that Al toxicity operates by causing a
depolymerization of both microtubules and microfilaments (Alfano et
al., 1993; Sasaki et al., 1997). Although these studies are consistent
with the hypothesis that the cytoskeleton could be a potential Al
target site, a correlation of the observed changes with those of the
temporal dynamics of growth inhibition has yet to be reported. Thus, an
assessment of whether the cytoskeleton is an initial symptom or a more
indirect or secondary symptom of Al exposure has not been possible.
Therefore, we conducted a detailed immunofluorescence study on the
effect of Al on the organization of microtubules and microfilaments in maize roots and correlated these effects with growth inhibition and
radial expansion of roots.
The data presented here indicate that toxic levels of Al result in a
significant cell-specific reorganization and stabilization of the
root's cytoskeleton, as summarized in Figure
10. The effect of Al on the
reorganization of the cytoskeleton was evident in the root-elongation
zone, consistent with results showing that this root zone is where Al
toxicity is first detectable (Ryan et al., 1993; Sivaguru and Horst,
1998). Generally, the effect of Al on the cytoskeleton was rapid,
coincided with the time course of growth inhibition, and was more
pronounced for microtubules than for microfilaments. In contrast to a
previous study showing that Al induced a depolymerization of cortical
microtubules in wheat roots (Sasaki et al., 1997), experiments
performed here with maize demonstrated that Al can induce a significant
stabilization of microtubules even in cells showing chronic symptoms of
Al stress.
View larger version (56K):
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| Figure 10.
Schematic diagram of a longitudinal section of a
maize root summarizing the changes in microtubules (MTs) (A) and
microfilaments (MFs) (B) in the elongation zone after exposure to Al.
A, Within 1 h of Al exposure, microtubules in the outer cortex
(oc) were stabilized in the transverse orientation. Oblique to random
microtubules were first detected in the inner cortex (ic) 3 h
after Al exposure and after 4 h in the stele (st). This shift in
microtubule orientation may lead to expansion of the inner cortical
cells, resulting in root swelling. By 12 h, outer cortical cells
with stabilized microtubules showed distorted shapes and were sloughed
off from the root, leading to lesions along the root surface. B,
Microfilaments were stabilized within 3 h after Al exposure.
Randomly oriented and highly bundled microfilaments in the inner cortex
and stele were detected 6 h after Al application. The
stabilization and reorganization of microfilaments occurred later than
that of microtubules.
|
|
Al toxicity is commonly associated with a swelling of the root apex
(Bennet et al., 1985; Ryan et al., 1993). Under the conditions used
here, radial expansion of the roots was first detected 6 h after
Al exposure and appeared to be caused by the abnormal isodiametric
expansion of cells located within the inner cortex (Fig. 2B). Our
observation of a selective, Al-induced microtubule reorientation in
these inner cortical cells, coupled with the widely proposed model for
microtubule orientation controlling cell expansion (Giddings and
Staehelin, 1991), may provide a mechanism to explain the Al-induced
inner cortical cell expansion leading to root swelling. However, it
still remains unclear from our studies whether the observed
reorientation of microtubules in the inner cortex is a direct effect of
Al or occurs in response to some secondary effector triggered by Al
stress.
Growth inhibition was detectable within 1 h (Fig. 1) and obvious
reorientation of microtubules in the inner cortex was detected after
3 h. Although Al can penetrate the outer two to three layers of
the root within 30 min (Lazof et al., 1994), most of it remains concentrated in the epidermal and outer cortical cells even after a
24-h exposure (Delhaize et al., 1993). Therefore, it is likely that the
disruption of microtubules in the inner cells could be an indirect
effect of Al. It is interesting to note that tissue-specific disruption
of microtubules in the inner cortex of maize roots has also been
reported to occur in ethylene-treated roots (Baluska et al., 1993a).
Studies on the relationship between Al toxicity and stress-related
ethylene synthesis (Reid, 1995) may provide important insight into the
mechanism of Al-induced microtubule reorientation and defense responses
(e.g. callose production) in inner cortical cells.
In contrast to the inner cortex, the overall orientation of
microtubules in the outer cortex and epidermis did not change upon Al
exposure. However, the microtubules in these tissues became increasingly stabilized and resistant to depolymerization after Al
application. This is similar to in vitro reports in mammalian systems
demonstrating that Al binding to microtubules and neurofilaments can
significantly delay their depolymerization (MacDonald et al., 1987;
MacDonald and Martin, 1988; Nixon et al., 1990; Shea et al., 1992). The
stabilizing effect of Al on the microtubules of cells on the outer
cortex was demonstrated by our auxin experiments, which showed that
even 1 h of Al exposure could block the IAA-induced reorientation
of microtubules from transverse to longitudinal arrays. Although the
data presented here cannot resolve the mechanisms by which Al prevented
auxin-induced microtubule reorientation, the stabilization caused by Al
was rapid (<1 h) and thus coincides with the inhibition of root
growth. The stabilizing effect of Al on the microtubules of the outer
cortex was also supported by other indirect observations, including:
(a) microtubules in Al-treated roots were less susceptible to
cold-induced microtubule depolymerization; (b) an increase in bundling
frequency in the microtubules of the outer cortex for both taxol and
Al-treated roots; and (c); taxol, like Al, prevented auxin-induced
microtubule reorientation.
The hypothesis that this stabilization of the microtubule network may
be partially responsible for inhibition of root growth is supported by
the very similar morphology of Al-treated roots with that of roots
treated with the microtubule-stabilizing drug taxol (see Fig. 6).
Furthermore, stabilizing microtubules in the transverse orientation
with taxol has also been shown to inhibit elongation growth (Baluska et
al., 1997; Weerdenburg and Seagull, 1988; this study). Therefore, in
addition to the orientation of microtubules (Giddings and Staehelin,
1991), it has been suggested that a dynamic microtubule network could
also play an important role in the control of elongation growth
(Weerdenburg and Seagull, 1988; Baskin et al., 1994).
Although taxol- and Al-treated roots exhibited similar morphology,
there were distinct differences in the onset of these effects. Radial
expansion in Al-treated roots was detected after 6 h, whereas for
taxol-treated roots expansion was detected only after 15 to 20 h.
Lesions in the surface of Al-treated roots were observed as early as
12 h after exposure, whereas lesions in taxol-treated roots were
observed only after 48 h (Fig. 6). Furthermore, microtubules in
the inner cortex of taxol-treated roots did not reorient but were
stabilized in the transverse orientation. These differences suggest
that a taxol-like stabilization of the microtubule network by Al may
not be singly responsible for the inhibition of root growth. In
addition to the effect of Al on the stability of microtubules in the
outer cortex, other growth-dependent processes that are independent of
microtubules may be simultaneously altered by Al exposure and thus
could account for the more pronounced and rapid effect of Al on the
growth and morphology of maize roots.
The concentration of Al chosen for this study (50 µM) was
the lowest that caused the maximal inhibitory effect on root growth rate (Fig. 1A) and the most pronounced effect on subsequent toxicity symptoms, such as swelling and the development of lesions (Figs. 2 and
6). However, concentrations lower than 50 µM also caused some growth inhibition (Fig. 1A) and root swelling (data not shown). Preliminary results showed that 12 h of exposure to 10 and 20 µM Al caused reorientation of microtubules in the inner
cortex (E.B. Blancaflor, D.L. Jones, and S. Gilroy, unpublished data). However, it is not known from this study whether the lower
concentrations of Al result in microtubule reorientation within the
same time frame as 50 µM Al or if at these lower Al
concentrations microtubules in the outer cortex are still stabilized.
Having established the patterns of cytoskeletal effects of an Al
concentration causing maximum effects on growth rate, it will be
important to determine the threshold Al concentration to elicit these
effects on the cytoskeleton.
Like microtubules, actin microfilaments were well preserved even after
long exposure to Al, but a qualitative assessment of these changes
proved more difficult than for microtubules because of their
less-ordered cellular arrangement, even in controls (see Fig. 8).
However, when Al-induced microfilament reorientation occurred, it was
very obvious and, as observed for microtubules, random arrays of
microfilaments were first detectable in the inner cortex. In contrast
to microtubules, however, this observed effect occurred after the onset
of growth inhibition and radial expansion (>6 h). Therefore, the
reorientation of microfilaments could simply be a consequence of the
changing polarity of the cells in the inner cortex. This is similar to
cytoskeletal changes accompanying wound healing in roots, wherein
microtubule reorientation has been shown to precede changes in the
polarity of cells, whereas the reorganization of microfilaments occurs
only after cells have already changed their growth direction (Hush and
Overall, 1992). However, it is possible that the fixation protocol used
in this study was not able to preserve the finer arrays of cortical
microfilaments in the elongating cells. The microfilaments in the
elongation zone observed in this study were primarily cytoplasmic
bundles and changes in the finer arrays of microfilaments may have been missed.
The experiments described above with the microfilament antagonist CB
also indicate that Al can induce a rapid ( 3 h) stabilization and
bundling of the actin network, in agreement with results presented for
soybean suspension cells (Grabski and Schindler, 1995; Grabski et al.,
1998). This observation is also supported by recent work showing that
genes encoding fimbrin, a known actin-bundling protein in animals, are
up-regulated in Al-treated wheat roots (Ortega et al., 1997). The
stabilizing effect of Al on both the microtubules and microfilaments
also indicates that both cytoskeletal components may respond to Al in a
coordinated way, as has been reported previously in taxol-treated rye
root tips (Chu et al., 1993).
Although these experiments indicate a significant interaction of Al
with the root's cytoskeleton, the exact mechanisms involved remain
unidentified. Because Al is known to rapidly enter cells (Lazof et al.,
1994), it is possible that it could interact directly with the
cytoskeletal elements, as demonstrated in vitro by MacDonald et al.
(1987). However, Al could also act indirectly through a modification of
the cell's physiochemical environment, signal transduction pathways,
or cytoskeletal anchors (Grabski et al., 1998). It is known that Al can
interact strongly with the plasma membrane, resulting in disruptions in
Ca2+ homeostasis, increased membrane rigidity,
and a blockage of lipid-mediated signal transduction cascades (Deleers
et al., 1986; Jones and Kochian, 1995, 1997). Because these are also
known to be integrally involved in the regulation of plant cytoskeletal
dynamics (Cyr, 1991; Xu et al., 1992; Shibaoka, 1994), it is likely
that Al may act both directly and indirectly on the root's
cytoskeleton.
| |
FOOTNOTES |
1
This work was supported by grants from the
National Science Foundation (to S.G.), the North Atlantic Treaty
Organization (to D.L.J. and S.G.), the Royal Society, and the Nuffield
Foundation (to D.L.J.).
2
These authors contributed equally to the paper.
*
Corresponding author; e-mail sxg12{at}email.psu.edu; fax
1-814-865-9131.
Received January 23, 1998;
accepted May 31, 1998.
| |
ABBREVIATIONS |
Abbreviations:
CB, cytochalasin B.
FDA, fluorescein
diacetate.
| |
ACKNOWLEDGMENTS |
We thank Jo Hughes for performing the microtome sectioning and
Richard Cyr for providing the anti-actin antibody. We also thank Sian
Ritchie and Richard Cyr for critical reading of the manuscript.
| |
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Abstract
Introduction