Early events responsible for aluminum toxicity ... - Wiley Online Library

Research

Early events responsible for aluminum toxicity symptoms in suspension-cultured tobacco cells Blackwell Publishing, Ltd.

Mayandi Sivaguru1, Yoko Yamamoto2, Zdenko Rengel3, Sung Ju Ahn2 and Hideaki Matsumoto2 1

Molecular Cytology Core Facility, 120, Life Sciences Center, University of Missouri-Columbia, Missouri 65211-7400, USA; 2Research Institute for

Bioresources, Okayama University, Chuo 2-20-1, Kurashiki 710-0046, Japan; 3Soil Science and Plant Nutrition, School of Earth and Geographical Sciences, The University of Western Australia, 35 Stirling Highway, Crawley WA 6009, Australia

Summary Author for correspondence: Hideaki Matsumoto Tel: +81 86 4341209 Fax: +81 86 4341249 Email: [email protected] Received: 18 May 2004 Accepted: 13 July 2004

• We investigated the aluminum (Al)-induced alterations in zeta potential, plasma membrane (PM) potential and intracellular calcium levels to elucidate their interaction with callose production induced by Al toxicity. • A noninvasive confocal laser microscopy has been used to analyse the live tobacco (Nicotiana tabacum) cell events by means of fluorescent probes Fluo-3 acetoxymethyl ester (intracellular calcium) and DiBAC4 (PM potential) as well as to monitor callose accumulation. • Log-phase cells showed no detectable changes in the PM potential during the first 30 min of Al treatment, but sustained large depolarization from 60 min onwards. Measurement of zeta potential confirmed the depolarization effect of Al, but the kinetics were different. The Al-treated cells showed a moderate increase in intracellular Ca2+ levels and callose production in 1 h, which coincided with the time course of PM depolarization. Compared with the Al treatment, cyclopiazonic acid, an inhibitor of endoplasmic reticulum Ca2+-ATPase, facilitated a higher increase in intracellular Ca2+ levels, but resulted in accumulation of only moderate levels of callose. Calcium channel modulators and Al induced similar levels of callose in the initial 1 h of treatment. • Callose production induced by Al toxicity is dependent on both depolarization of the PM and an increase in intracellular Ca2+ levels. Key words: Al toxicity, callose formation, intracellular calcium, Nicotiana tabacum, plasma membrane depolarization, zeta potential. New Phytologist (2005) 165: 99–109 © New Phytologist (2004) doi: 10.1111/j.1469-8137.2004.01219.x

Introduction Although substantial literature exists on aluminum (Al) toxicity in plants, the specific mechanism by which Al inhibits plant growth and development is elusive (Horst, 1995; Kochian, 1995; Matsumoto, 2000). Effects of Al on the potential across the plasma membrane (PM) (transmembrane potential) are the early response to Al toxicity in various experimental systems (Ahn et al., 2001, 2004). Depolarization of the transmembrane potential was observed in experiments with intact roots of Beta vulgaris (Lindberg et al., 1991) or PM-enriched microsomes (Siegel & Haug, 1983) at relatively high pH values (e.g. 6.5).

www.newphytologist.org

In addition, at pH 4.0 or 5.0, Al caused hyper-polarization of the PM of intact Beta vulgaris roots (Lindberg et al., 1991). Aluminum caused instantaneous PM depolarization in root cells of an Al-sensitive maize cultivar, with the intensity of depolarization varying for different growth zones (Sivaguru et al., 1999a). In intact Triticum aestivum roots, small Al-related depolarization was found in the Al-sensitive cultivar Scout but not in the Al-tolerant cultivar Atlas (Miyasaka et al., 1989), whereas no significant effect of Al on the PM potential was found in the follow-up study on the same cultivars (Huang et al., 1992). However, the zeta potential (i.e. membrane surface potential)

99

100 Research

of PM vesicles isolated from roots of cv. Scout was 26% more negative than that of vesicles from cv. Atlas, allowing more Al to bind to the Scout vesicles (Yermiyahu et al., 1997), and causing greater Al toxicity. In line with these observations, greater depolarization of zeta potential was observed recently in vesicles isolated from Al-sensitive ES8 near-isogenic wheat line compared with the Al-tolerant ET8 (Ahn et al., 2004). Such a result indicates that zeta potential may be a more appropriate factor related to Al toxicity than the measurement of transmembrane potential. Hence, the study presented here compared the effects of Al on both the transmembrane and zeta potentials. Interactions between Al and Ca2+ have long been implicated in Al toxicity in plants (Rengel, 1992a, 1992b, 1996; Ryan & Kochian, 1993), with Al disrupting Ca2+-dependent metabolic processes, including cell division and elongation. Studies using Ca2+-sensitive fluorescent dyes and Ca2+ imaging techniques have detected Al-induced changes in cytosolic Ca2+ levels in several plant systems (for a review see Rengel & Zhang, 2003). Most studies demonstrated an increase in cytosolic Ca2+ activity as a consequence of Al toxicity, for example in intact wheat (Zhang et al., 1998; Zhang & Rengel, 1999) and rye roots (Ma et al., 2002), excised barley roots (Nichol & Oliveira, 1995), Arabidopsis root hairs ( Jones et al., 1998) and protoplasts isolated from wheat roots (Lindberg & Strid, 1997). Accumulation of callose is one of the most frequently used markers of Al toxicity (Zhang et al., 1994; Wissemeier & Horst, 1995). To date, no study simultaneously investigated the mechanism of callose formation and dynamics of intracellular Ca2+ and the PM potentials (surface and transmembrane). However, Bhuja et al. (2004) studied intracellular Ca2+ and callose formation together and concluded that an increase in intracellular Ca2+ (albeit required for initiating synthesis of callose; see Kauss, 1996) was not the only factor responsible for callose production in the presence of Al. The present study highlights the complex and intrinsic relationships leading to callose accumulation under Al toxicity by studying the role of intracellular Ca2+ as well as the surface and transmembrane potentials of intact tobacco cells.

Materials and Methods Cell culture and experimental conditions A nonchlorophyllous tobacco cell line SL derived from Nicotiana tabacum L. cv. Samsun (Nakamura et al., 1988) was used. Cells grew in a modified version of Murashige-Skoog’s (MS) medium (Yamamoto et al., 1994, 1997) at pH 5.0 (measured after autoclaving). Cells were serially subcultured at 7-d intervals at a 1 : 15 dilution. During subculture, cells exhibited a logarithmic phase of growth from days 2–5 and reached the stationary phase on day 7. For the Al treatment, cells in the logarithmic phase (day 4) were washed twice with

New Phytologist (2005) 165: 99–109

a simplified MS medium containing only 3 m CaCl2 and 3% (w : v) sucrose (pH 5.0) (referred to hereafter as Ca medium) and resuspended in the Ca medium (pH 4.5) at the cell density of 100 mg fresh weight per 10 ml as described previously (Ikegawa et al., 2000). Resuspending the cells at this density enabled us to use a low ionic strength medium, where Al was available predominantly as monomeric ion (Al3+). Determination of zeta potential in protoplasts using light scattering electrophoresis Control and Al-treated protoplasts, isolated using the standard protocol (Ahn et al., 2001), were used to measure zeta potential by means of free-flow electrophoresis (using an ELS-8000 apparatus; Otsuka Electronics, Osaka, Japan). The electrophoresis medium (chamber buffer) was presterilized by passing through a glass fiber filter followed by a membrane filter (0.45 µM). Cells were washed several times in the chamber buffer and kept in it for at least 30 min prior to measurement. Freshly isolated protoplasts (50 mg cells per 5 ml) were diluted 10 times with the same buffer; the sample (equal volume basis) was then added to an acrylic electrophoresis cell. A computer connected to the detection system was used to perform measurements. After each measurement, cells were washed several times in the chamber buffer before new measurement commenced. The zeta potential was determined against Latex (Dow Chemical, Japan) as a standard. The zeta potential (ζ) of PM vesicles was calculated from the electrophoretic mobility (µ) using the Smoluchowski equation: ζ = µη/εε0 (η is the viscosity, ε is the relative dielectric constants of the buffer and ε0 is the dielectric constant of vacuum) (Gimmler et al., 1991). Protoplasts were isolated from two individual experiments having three replicates for each treatment and time point. Quantitative determination of Al Aluminum content was quantified using atomic absorption spectrometry (Hitachi 3000; Hitachi Tokyo, Japan) after incubating cells in 2  HCl for 48 h following experimental treatments. Two independent experiments comprising three replicate sets of samples at each time point were analysed. Fluorescent probes The Ca2+-sensitive fluorescent dye acetoxymethyl ester Fluo3 (cell-permeant) for imaging live-cell intracellular Ca2+ levels, and the potentiometric probe DiBAC4, an anionic oxonol, suitable for a noninvasive monitoring of the PM potential, were purchased from Molecular Probes (Eugene, OR, USA). The Ca2+ indicator did not interact with Al

www.newphytologist.org © New Phytologist (2005)

Research

(Zhang et al., 1998). This calcium indicator has been used successfully with Al in intact wheat (Zhang et al., 1998; Zhang & Rengel, 1999; Bhuja et al., 2004) and rye roots (Ma et al., 2002) as well as in Chamelaucium uncinatum pollen tubes (Zhang et al., 1999). The membrane potential dye DiBAC4 cited as ‘bis-oxonol’ has been extensively used in animal cells as well as in few plant experiments (Britten et al., 1992; Johannes et al., 1992; Pouliquin et al., 1999). Loading of Fluo-3/AM and DiBAC4 into tobacco cells Log-phase cells were incubated with the appropriate dye using the method described by Zhang et al. (1998). Briefly, after two washes in Ca medium (pH 5.0) the cells were incubated in a medium containing 20 µ Fluo-3/AM, 50 m sorbitol (pH 4.3) at 4°C for 90 min in the dark, followed by 90-min incubation in the aerated Ca-medium with 50 m sorbitol at 20°C in the dark. The low-temperature incubation inhibits the extracellular hydrolysis of the dye, which would have prevented the diffusion of Fluo-3/AM across the PM. The intact Fluo-3/AM that diffused into the cells was hydrolysed by intracellular esterases at 20°C, releasing Ca2+-reporting, membrane-impermeable dye inside the cells (Zhang et al., 1998). The loaded cells were protected from light during the experimental period. In general, the potentiometric dye DiBAC4 was loaded into tobacco cells in the same way as Fluo-3/AM (low temperature coupled with the low-pH Ca-medium containing 50 m sorbitol). DiBAC4 belongs to a class of slow but stableresponse probes, which enter depolarized cells quickly, bind to intracellular membranes and exhibit enhanced fluorescence or red spectral shifts (Montero et al., 1994). By contrast, subsequent return of the PM potential to resting levels or even hyperpolarized conditions results in extrusion of the dye and thus a decrease in fluorescence intensity. Incubating the dyeloaded tobacco cells in 100 m KCl tested this property. Within 15–30 min, cells showed a marked increase in red fluorescence along the cytoplasmic periphery, indicating PM depolarization (data not shown). Although the manufacturer’s protocol suggests an increase in fluorescence intensity of around 1% for each 1 mV depolarization, we did not attempt to quantify the change in the PM potential in arbitrary units. Instead, we have quantified the zeta potential, an indicator that yielded qualitative results comparable to those obtained in the experiments with DiBAC4. Treatments with Al and the Ca2+-channel modulators After loading, the cells were monitored under the microscope for viability and labeling uniformity. Cytoplasmic streaming was used as a marker for viability. Cells or cell clusters with uniform fluorescence, especially around the nuclei and cytoplasmic periphery, constituted approximately 94% of all the cells; they were used for imaging experiments. Loaded

© New Phytologist (2005) www.newphytologist.org

cells were suspended at a density of 100 mg 10 ml−1 in the treatment solution with Al (AlCl3.6H2O, a range of concentrations), or the Ca2+ ionophore A-23187 (20 µ), or the inhibitor of endoplasmic reticulum Ca2+-ATPase cyclopiazonic acid (CPA, 20 µ) from 15 min to a maximum of 12 h. During the treatments, cells were kept on a shaker operating at 100 r.p.m. in the dark. After the end of the specified period, several aliquots of cells were sampled and the images were obtained instantly. When the treatments were performed directly under the microscope using the flow-cytometry method, the loaded cells were suspended in poly -lysinecoated delta T-3 dishes (Bioptechs, Butler, PA, USA) with No. 1 cover slips as the bottom. Experiments testing Al and calcium channel modulator effects on intracellular Ca2+ levels and PM potentials were repeated twice, each time with three to four replicates for each time-point or concentration. The images presented are the representative patterns for a particular treatment across the two experiments. Image acquisition and evaluation of intracellular Ca2+, PM potential and callose using live-cell confocal laser scanning microscopy Fluorescence images of intracellular Ca2+, membrane potentials and callose were recorded under the Bio-Rad MRC 600 (Bio-Rad, San Diego, CA, USA) (for intracellular Ca2+) and Zeiss LSM 510 (Zeiss, Oberkochen, Germany) (membrane potentials and callose) microscopes using ×40 (1.4 NA) and ×100 (1.3 NA) Plan-Apochromat objectives. Fluo-3/AM fluorescence was monitored using standard optical filters for fluorescein excitation (488 n) and emission (515 n) wavelengths. Callose formation was assessed by monitoring an increase in autofluorescence at the 454-n wavelength of Argon laser or under UV filters. In some experiments, simultaneous analysis of callose accumulation and the PM potential in intact cells was performed using 488 n excitation wavelength followed by superimposing images to assess whether the callose formation occurred only in depolarized cells. Images were mostly obtained through the median cell plane and were either left with original color or pseudocolored for easy assessment of intensity gradients. In the confocal laser scanning microscope (CLSM) reuse mode, constant scanning parameters such as pinhole, amplifier offset and gain were employed for a respective dye, which allowed intensity comparisons between treatments. In the case of intracellular Ca2+ and the PM potential, each frame consisted of 4–6 individual cells (under ×100 magnification). A total of more than 30–55 frames were recorded for each treatment. The frequency (%) of a particular cell type/pattern was calculated by projecting all these frames. The percentage of a particular cell type was independently calculated based on the intensity change from each treatment/time-point for two


101

102 Research

experiments and averaged. These numbers were presented separately in a table for easy assessment of the response pattern. Images recorded on Fuji Provia-Professional (100 ASA) slide film were assembled with Adobe Photoshop 7.0 J (Adobe System, San Jose, CA, USA) and printed using a Fujix Pictrography 3000 digital printer. Determination of callose After treatments with Al and the Ca2+ ionophores, the callose contents in the log- and the stationary-phase cells (cells in 10 ml culture, corresponding to 100 mg fresh weight at the start of the treatment) were quantified using decolorized aniline blue technique (Kauss, 1996) and expressed as Curdlan equivalents. Two independent experiments were performed with four replicate treatments for each Al/Ca2+ ionophore combination and for each time point.

Fig. 1 The time-course of Al effects on the zeta potential of tobacco (Nicotiana tabacum) protoplasts isolated from the log-phase cells. Note a clear depolarization in response to Al and a moderate recovery in Al-free medium for 3 h following Al treatment for 6 h and 12 h. Data are means ± SE of three independent replicates and representative of two individual experiments. Circles, control; squares, +50 µM Al; triangles, after 3 h wash in Al-free medium.

Statistics All experiments reported here were repeated at least twice, with three to four individual replicates to reduce variation. Where necessary, Student’s t-test was performed in Sigma Plot version 3.0 to separate means of data points at 95% confidence levels.

Results The growth response of tobacco cells to Al was similar to that observed in previous studies using the same material under identical growth conditions (Sivaguru et al., 1999b). An increase in growth (as determined by fresh weight) of logphase tobacco cells was abolished almost completely (reaching only 30% of control) after 12 h of exposure to Al (Sivaguru et al., 1999b). Impact of Al on zeta potential The zeta potential of tobacco protoplasts was measured because it might be an early response to Al preceding PM depolarization. Exposure of cells to Al for 1 h resulted in no change in zeta potential (Fig. 1). However, significant depolarization of zeta potential was detected in cells treated with Al for 6 h and especially 12 h. There was no change in zeta potential after the 12-h period in the absence of Al. The zeta potential of cells treated with Al for 6 or 12 h recovered moderately upon transfer to the Al-free solution for 3 h (Fig. 1), suggesting a large degree of irreversible Al binding to the PM. Al accumulation Accumulation of Al showed a saturable response over time, with a steep initial increase after 1 h followed by a much


Fig. 2 An Al-accumulation pattern in log-phase tobacco (Nicotiana tabacum) cells subjected to 50 µM Al for various lengths of time. Data are means ± SE of three independent replicates and representative of two individual experiments.

slower increase (or even leveling off ) after 6 h and 12 h of exposure to Al (Fig. 2). Impact of Al on PM potential To our knowledge, the effect of Al on the PM potential has not been shown in intact suspension-cultured cells. The conventional method of measuring transmembrane potential by impalement in the case of suspension cultured cells might yield results for one particular cell at a time, but cannot provide a measure of a potential impact on neighboring cells. Therefore, we have exploited a technique of intact cell imaging of the PM potential using a fluorescent dye/marker. During the first 30 min of exposure to Al, there was no discernible difference on the PM potential of cells preloaded with the anionic DiBAC4 (oxonol) dye (Fig. 3A,B). However, large PM depolarization was detected from 60 min onwards, persisting for the extended period (720 min) (Fig. 3C–F).


Research

Fig. 3 Confocal pseudo-color images showing Al-mediated alterations in plasma membrane (PM) potential in the log-phase tobacco (Nicotiana tabacum) cells. Images were obtained at the beginning of the experiment at time = 0 (A) followed by subjecting the same batch of cells to 50 µM Al in the calcium (Ca) medium containing sorbitol (50 mM). The images were obtained at the time-points indicated (in min) (B–F). Separate batches of cells treated with Al for 360 min (G) and 720 min (H) were transferred to Ca medium without Al and allowed to recover for a period of 180 min. Parallel control cells were shown from the same batch treated without Al for 720 min (I). The blue color is the background and the deep red color indicates the maximum PM depolarization, as indicated in legend (J). Representative images are from at least two independent experiments each consisting of three replicates. Bar, 40 µM.

The cells that had been exposed to Al for 360 min showed marked reversal of the Al effect on the PM potential during the subsequent 180-min recovery period without Al ( Fig. 3G). However, the cells treated with Al for 720 min appeared to have sustained irreversible PM depolarization (Fig. 3H). Cells kept under identical conditions without Al for 720 min (controls) showed no depolarization (Fig. 3I).

upon increasing duration of treatment (Fig. 4G). Approximately one-quarter of cells showed highest response (35%) and half of them (54%) moderately responded to 100 µ Al during 60-min exposure. These numbers reversed upon prolonged time at the same Al concentration (i.e. 95% of cells showed the highest and 3% of cells a moderate response; Fig. 4F,G and Table 1). The control (–Al) cells loaded with the dye showed no response even after 300 min (Fig. 4C).

Impact of Al on intracellular calcium levels In the absence of definitive reports on the cause–effect relationship between PM depolarization and the Ca2+ influx, this study presumed that the opening of calcium channels or increase in intracellular calcium causes PM depolarization (see Fig. 1 in Dennison & Spalding, 2000). This presumption is also based on an earlier report by Sivaguru et al. (2003), where pretreatments with gadolinium or lanthanum (Ca2+ channel blockers) abolished the Al-induced PM depolarization. Around 88–100% of cells incubated in 50 µ Al for 60 min (see Fig. 4D,E and Table 1) showed a moderate increase in the intracellular Ca2+ compared with the loaded control (Fig. 4B). Cells treated with 100 µ Al showed a moderate increase (60 min, Fig. 4F) in the intracellular Ca2+ levels at the cytoplasmic periphery, and this pattern intensified


Impact of calcium ionophores on the intracellular calcium levels The pattern of changes in the intracellular Ca2+ in tobacco cells (Fig. 4) caused by Al (an increase in the intracellular Ca2+, especially at the cytoplasmic periphery and around nuclei) was similar to the pattern recorded in the cells treated with the Ca2+-channel modulator A-23187 (Fig. 4H) or inhibitor of the endoplasmic reticulum Ca2+-ATPase, CPA (Fig. 4I). The cells were sampled at approximately 90 min after commencement of treatments. For A-23187, 35% of cells showed the highest response and 60% showed a moderate response, in accordance with results obtained using 100 µ Al for 60 min (Fig. 4F and Table 1). In the case of CPA (90 min), the response was equivalent to 100 µ Al for


103

104 Research

Fig. 4 Confocal pseudo-color images showing alterations in intracellular Ca2+ levels in response to Al or Ca2+ channel modulators in tobacco (Nicotiana tabacum) log-phase cells. Control cells were either not loaded (A) or loaded with the Ca2+ indicator Fluo-3/AM ester (B). Aluminum (D–G) or Ca2+ channel modulators A-23187 (H) and CPA (I) caused alterations in intracellular Ca2+ levels. Two frames (D–E) of 50 µM Al were shown for uniformity in response. Cells left in –Al medium for 320 min and handled in the same way as the treatments did not show an increase in intracellular Ca2+ levels (C). When the Al concentration was increased to 100 µM (F,G), the response (increase in the Ca2+ signal at the cytoplasmic periphery) was similar to the effect caused by Ca2+-channel modulators (H,I). The color code is black as background and magenta as the highest Ca2+ level. The response pattern has been divided in four major classes (indicated as asterisks) and a representative cell for a given class indicated on the images (see Table 1 for details of analysis). The images are representative of two independent experiments, each having three to four replicates. Bar in (A), 25 µM for all images. Table 1 Analysis of the effect of Al- and Ca2+-channel modulators A-23187 and cyclopiazonic acid (CPA) on the intracellular Ca2+ levels in logphase tobacco (Nicotiana tabacum)1

Calcium response pattern2

Controls A

B

C

Al 50 µM D

E

Al 100 µM F

G

A-23187 H

CPA I

(***) Highest response (**) Moderate response (*) No response (O) Not determined

– – – –

– – 93 7

– – 100 –

– 88a 10 2

– 100a – –

35a 54b 7 4

95b 3c 2 –

35a 60b – 5

82b 15d – 3

1 A–I are images from Fig. 4 and values are percentages. Data reported have 3–6% of error across the two independent experiments each having three or four replicates. Means with different letters across the treatments are significantly different at P = 0.05 (Student’s t-test). Cells that did not fit in any patterns were grouped as ‘(O) Not determined’. 2 Grading of the response is based on the cells expressing increased cytoplasmic Ca2+ in response to treatment and for other details see the Materials and Methods section.

310 min (Fig. 4G and Table 1), i.e. 82% of cells showed the highest response and 15% reacting moderately to CPA (Table 1). These results indicate the calcium ionophores are faster and more effective in increasing the intracellular Ca2+ levels than Al in tobacco cells.

observed after 6 h of exposure to Al (50 µ). Optical sections performed over the surface of cells revealed a typical ‘spotty’ appearance of callose (Fig. 5D,E). The control (–Al) cells kept under identical conditions as the Al-treated cells for 720 min did not show callose accumulation (Fig. 5F).

Visualizing the Al-induced callose accumulation

Simultaneous visualization of calcium-ionophoreinduced changes in intracellular calcium and callose production in intact cells

The accumulation of callose in the log-phase cells increased with the duration of exposure to Al, as visualized by an increase in fluorescence intensity (Fig. 5). Al-induced callose was detectable only after 3 h ( Fig. 5B), with substantial levels


Using a dual detection technique for the first time that (1) exploits an increase in the autofluorescence of cell walls due to


Research

Fig. 5 Confocal images showing Al-induced (50 µM) callose formation in log-phase tobacco (Nicotiana tabacum) cells at indicated time points. (A–C,F) A single optical section through the median plane; (D,E) optical sections (7 µM) from the cell surface towards the center overlaid to clarify the callosic spots. Images are from two independent experiments, each having three replicates. Bar, 40 µM (E), 20 µM (A–D,F).

increased callose production, and (2) the use of the PM potential indicator, we could simultaneously monitor these two parameters in intact cells (Fig. 6). Even though preliminary experiments revealed significant changes in the intracellular Ca2+ levels after 90 min, we increased the exposure time to 3 h to enhance the differences in the response between these two markers. In the control cells, there was no callose accumulation (Fig. 6A1), and the PM showed only resting potential levels (Fig. 6A2). Therefore, the merged image (Fig. 6A3) showed no yellow coloration because there was no callose (Fig. 6A3). However, after the Ca2+ ionophore treatment (Fig. 6, sets B and C), enhanced callose production and the PM depolarization were apparent from the increased yellowto-green ratio in the merged images (Fig. 6, B3 and C3). As a positive control, cells incubated with CPA but without the marker for the PM potential accumulated callose to a similar extent as cells loaded with the marker for the PM potential


Fig. 6 Confocal images of intact cells showing simultaneous analysis of PM potential and callose formation in response to Ca2+ channel modulators. Log-phase tobacco (Nicotiana tabacum) cells were treated with Ca2+-channel modulators A-23187 and CPA (each 20 µM) for 3 h. An increase in the red fluorescence indicates higher callose accumulation, whereas an increase in green fluorescence shows plasma membrane (PM) depolarization. Only when these two phenomena coincide do the red and green pixels merge and form yellow pixels. Note there was neither callose accumulation (A1) nor plasma membrane depolarization in controls (set A); hence, there was no yellow component in the merged image (A3). Nearly all pixels were colocalized to form yellow pixels in treatments (B3 and C3). A positive control has been used for the CPA treatments (no PM potential dye was loaded), showing the callose signal (D1 and D3). Representative images are shown from two independent experiments, each having three replicates. Bar, 20 µM.

(Fig. 6, set D). Only the CPA-induced callose showed the distribution pattern similar to Al-induced ‘spotty’ callose accumulation (Fig. 6, C1, C3, D1 and D3). Quantification of callose induced by Al or calcium ionophores We hypothesized that both Al and calcium channel modulators could induce callose production within an hour. However, in the case of Al we could observe the callose accumulation using the confocal microscopy only after 3 h (Fig. 5B). Hence, we used the highly sensitive, spectrophotometric quantification


105

106 Research

Fig. 7 The effect of Al (50 or 100 µM) and Ca2+-channel modulators A-23187 and CPA (at 20 µM) on the callose formation at three time points (open columns, 1 h; tinted columns, 6 h; closed columns, 12 h) in log-phase tobacco (Nicotiana tabacum) cells. Background or control callose levels were subtracted from each treatment. Means with different letters across the treatments are significantly different at P = 0.05 (Student’s t-test). Data are means ± SE.

method of callose (see Wissemeier et al., 1987). As expected, 1-h treatments with either Al or calcium ionophores showed similar (and significant) accumulation of callose (Fig. 7). Although the ionophores were quicker than Al in increasing the intracellular Ca2+ levels (Fig. 4), they did not produce as high callose accumulation as Al over prolonged time (Fig. 7). Even though CPA showed similar accumulation of callose when assessed by microscopy (Fig. 6, C1 and C3), and increased intracellular Ca2+ in 1.5 h to a similar extent as 100 µ Al after 5 h (Fig. 4G,I), CPA did not induce accumulation of callose to the same level as 100 µ Al when the quantitative method was used for measuring callose (Fig. 7).

Discussion Although several mechanisms were studied with the aim of resolving the toxic effects of Al (Rengel & Zhang, 2003), so far the early events associated with Al toxicity such as zeta potential, transmembrane potential, intracellular Ca2+ levels and callose formation have not been analysed simultaneously in any plant system. As indicated in the introduction, two of these components have been investigated simultaneously in a study (Bhuja et al., 2004) published while this manuscript was under review. In addition, for the first time in Al studies, a fluorescent PM potential indicator was used, allowing acquisition of data from a large number of cells concurrently. This would be a promising advantage over impaling the intact cells because impalement itself could trigger several signal transduction cascades and callose production. At present, investigating early events in Al toxicity (on the scale of seconds and finer) is technically not feasible. Even though we studied phenomena involved in cellular signal transduction cascades (PM potential and intracellular Ca2+


levels), the relatively prolonged kinetics of events recorded precludes statements about signaling. Changes in the zeta potential are one of the early responses to Al toxicity (Miyasaka et al., 1989; Yermiyahu et al., 1997), possibly owing to Al binding onto the PM. The differences in the magnitude of negative charges on the surface of the plasma membrane are expected to differentially attract the positively charged Al ions (Yermiyahu et al., 1997; Ahn et al., 2004), which may alter phospholipid profile thereby affecting lipid-mediated signaling (Kochian, 1995; Matsumoto, 2000). Such alterations of electrical properties of the membrane surface may destabilize the PM. Our results indicate that depolarization of tobacco cells (observed in terms of both zeta potential and the PM potential) was comparable to depolarization of PM in response to Al in actively growing root tips of the Al-sensitive maize (Sivaguru et al., 1999a) or Arabidopsis (Sivaguru et al., 2003). Similarly, a recent report on two near-isogenic wheat lines differing in sensitivity to Al showed depolarization of zeta potential only in the Al-sensitive line (Ahn et al., 2004). Log-phase tobacco cells accumulated significant amounts of Al after 1 h (Fig. 2). Without being able to distinguish between apoplastic and symplastic location of accumulated Al (Taylor et al., 2000), we assume that a portion of Al measured was located inside the cells, making it impossible to rule out the possibility of intracellular lesion causing, or at least contributing to, Al toxicity. In line with this proposal, Vitorello and Haug (1997) showed a rapid (< 30 min) internalization of Al as Al–morin complex in BY-2 tobacco suspension cells. In our study, a large increase in Al content was measured within 1 h of exposure (Fig. 2), depolarization of zeta potential was not observed at this time (Fig. 1), and irreversible damage to cells (in terms of PM depolarization) occurred only after 12 h (Fig. 3). Such results suggest that Al internalization may be a slow process, with most Al initially accumulating in the cell wall (see Tabuchi & Matsumoto, 2001). Aluminum increases intracellular Ca2+ in several plant systems (see Introduction). In the present study, Al also increased the intracellular Ca2+ levels (Fig. 4, Table 1). This level of increase was lower than that achieved by the two Ca2+ ionophores (Fig. 4). Sustained (up to 1 h) Al-related increases in intracellular Ca2+ levels in tobacco cells were correlated with decline in growth (Vitorello & Haug, 1996). Also, oxidative stress increased intracellular Ca2+ levels in tobacco (Price et al., 1994), and Al has been shown to induce genes involved in oxidative stress (Ezaki et al., 2000). An increase in intracellular Ca2+ may have an important role in the expression of Al toxicity because increased intracellular Ca2+ would eliminate transient spikes in intracellular Ca2+ activity caused by environmental signals, including activation of ionotropic glutamate receptors (Dennison & Spalding, 2000), whereby PM depolarization kinetics closely followed a rapid spike in intracellular Ca2+ levels. Aluminium toxicity triggered accumulation of callose in log-phase tobacco cells as early as 1 h after exposure (Fig. 7),


Research

a phenomenon closely linked with Al toxicity in various experimental systems (Zhang et al., 1994; Wissemeier & Horst, 1995; Sivaguru & Horst, 1998; Sivaguru et al., 1999a,b, 2000a,b). An increase in intracellular Ca2+ is a prerequisite for the synthesis of callose in plant cells (Kauss, 1996). Given the spotty pattern of callose accumulation after exposure to Al (Fig. 5) or CPA (Fig. 6), it is possible that localized increases in intracellular Ca2+ (Fig. 4) can occur in the areas rich in endoplasmic reticulum around secondary plasmodesmata or in adjacent areas in these suspension-cultured cells. Even though we do not provide direct evidence for such claim, spotty distribution of viral movement proteins have been found to colocalize with callose in protoplasts isolated from cultured cells (Monzer, 1991; Heinlein et al., 1998; Itaya et al., 1998). We have observed similar spotty callose production in response to Al in protoplasts isolated from suspension cultured maize cells (data not shown). In intact plant cells it has been well documented that the callose production occurs at these discrete sites, implicating these sites as Altoxicity lesions (Holdaway-Clarke et al., 2000; Sivaguru et al., 2000a). The other possibility is localization of callose where protein aggregates of cellulose synthase occur; cellulose synthase may be involved in producing callose (Delmer, 1999) in the presence of Al (Teraoka et al., 2002), with callosic deposits being higher around those protein aggregates. Such a change between callose and cellulose production may be involved in an adaptive response to Al toxicity because most Al-tolerant plants do not synthesize callose as much as sensitive plants (Llugany et al., 1994), thus maintaining a high rate of cellulose synthesis and hence a good growth rate. The callose induction could not be visualized within the 1-h time frame using the confocal microscope, suggesting quantitative analysis of callose using a fluorescence spectrophotometer should also be performed to corroborate the confocal microscopy results. A-23187 (a Ca2+ ionophore) did not induce callose pattern that was similar to the Al-induced one (Figs 5 and 6), suggesting the source of Ca2+ responsible for callose production might be from the internal sources. Similar observations were made by Bhuja et al. (2004), where A23187 was very effective in increasing the intracellular Ca2+ but showed only an 11-fold increase in callose levels compared with the 30-fold increase with Al. In the study presented here, the rapid Al accumulation (Fig. 2), the Al-induced increase in intracellular Ca2+ (Fig. 4), and the PM depolarization (Fig. 3) were all observed within the same time-frame (1 h), suggesting that callose production under Al toxicity requires PM depolarization and an increase in intracellular Ca2+. Nevertheless, we cannot rule out the possibility that other factors might also contribute to sustained callose production under prolonged Al treatments. Cells produce and deposit the callose at the external face of the PM to protect it from the toxic effects of Al, which is a response similar to the one occurring during pathogen infection. In the light of these points, it would be intriguing to analyse


the Al effects in mutant plants which lack the capacity to synthesis callose. Investigating the localization and distribution of PM-embedded Ca2+ channels and other potential Alreceptors on the surface of the cell membrane may extend our understanding of signaling cascade in Al toxicity as well as of interactions between PM potential, intracellular Ca2+ and callose accumulation.

Acknowledgements This work was financially supported by the Program for Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN), Grant-in-Aid for Scientific Research (grant no. 14206008) from the Ministry of Education, Culture, Sports, Science and Technology of Japan, Ohara Foundation for Agricultural Sciences and the Japan–Australia research cooperative program from Japan Society for the Promotion of Science to H. M., Australian Research Council International Linkage grant to Z. R. and a postdoctoral fellowship from the Japan Society for the Promotion of Science to M. S. We thank Dr Walter Gassmann and Sharon Pike, University of Missouri-Columbia, for their constructive critical comments on the manuscript.

References Ahn SJ, Sivaguru M, Osawa H, Chung GC, Matsumoto H. 2001. Aluminum inhibits the H+-ATPase activity by permanently altering the plasma membrane surface potentials in squash roots. Plant Physiology 126: 1381–1390. Ahn SJ, Rengel Z, Matsumoto H. 2004. Aluminum induced plasma membrane surface potential and H+-ATPase activity in near isogenic wheat lines differing in tolerance to aluminum. New Phytologist 162: 71–79. Bhuja P, McLachlan K, Stephens J, Taylor G. 2004. Accumulation of 1,3β--glucans, in response to aluminum and cytosolic calcium in Triticum aestivum. Plant Cell Physiology 45: 543–549. Britten CJ, Zhen RJ, Kim EJ, Rea PA. 1992. Reconstitution of transport function of vacuolar H(+)-translocating inorganic pyrophosphatase. Journal of Biological Chemistry 267: 21850–21855. Delmer DP. 1999. Cellulose biosynthesis: exciting times for a difficult field of study. Annual Review of Plant Physiology and Plant Molecular Biology 50: 245–276. Dennison KL, Spalding EP. 2000. Glutamate-gated calcium fluxes in Arabidopsis. Plant Physiology 124: 1511–1514. Ezaki B, Gardner RC, Ezaki Y, Matsumoto H. 2000. Expression of aluminum-induced genes in transgenic Arabidopsis plants can ameliorate aluminum stress and/or oxidative stress. Plant Physiology 122: 657–665. Gimmler H, Treffny B, Kowalski M, Zimmermann U. 1991. The resistance of Dunaliella acidophila against heavy metals: The importance of the zeta potential. Journal of Plant Physiology 138: 708–716. Heinlein M, Padgett HS, Scott Gens J, Pickard BG, Casper SJ, Epel BL, Beachy. RN. 1998. Changing patterns of localization of the tobacco mosaic virus movement protein and replicase to the endoplasmic reticulum and microtubules during infection. Plant Cell 10: 1107–1120. Holdaway-Clarke TL, Walker NA, Hepler PK, Overall RL. 2000. Physiological elevations in cytoplasmic free calcium by cold or ion injection result in transient closure of higher plant plasmodesmata. Planta 210: 329–335.


107

108 Research Horst WJ. 1995. The role of the apoplast in aluminum toxicity and resistance of higher plants: a review. Journal of Plant Nutrition and Soil Science 158: 419–428. Huang JW, Shaff JE, Grunes DL, Kochian LV. 1992. Aluminum effects on calcium fluxes at the root apex of aluminum-tolerant and aluminumsensitive wheat cultivars. Plant Physiology 98: 230 –237. Ikegawa H, Yamamoto Y, Matsumoto H. 2000. Responses to aluminum of suspension-cultured tobacco cells in a simple calcium solution. Soil Science and Plant Nutrition 46: 503 –514. Itaya A, Woo YM, Masuta C, Bao Y, Nelson RS, Ding B. 1998. Developmental regulation of intercellular protein trafficking through plasmodesmata in tobacco leaf epidermis. Plant Physiology 118: 373–385. Johannes E, Brosnan JM, Sanders D. 1992. Parallel pathways for intracellular Ca2+ release from the vacuole of higher plants. Plant Journal 2: 97–105. Jones DL, Gilroy S, Larsen PB, Howell SH, Kochian LV. 1998. Effect of aluminum on cytoplasmic Ca2+ homeostasis in root hairs of Arabidopsis thaliana (L.). Planta 206: 378 –387. Kauss H. 1996. Callose synthesis. In: Smallwood M, Knox JP, Bowles DJ, eds. Membranes: specialized functions in plants. London, UK: Bios Scientific Publishers, 77–92. Kochian LV. 1995. Cellular mechanisms of aluminum toxicity and resistance in plants. Annual Review of Plant Physiology and Plant Molecular Biology 46: 237–260. Lindberg S, Strid H. 1997. Aluminium induces rapid changes in cytosolic pH and free calcium and potassium concentrations in root protoplasts of wheat (Triticum aestivum). Physiologia Plantarum 99: 405 – 414. Lindberg S, Szynkier K, Greger M. 1991. Aluminium effects on transmembrane potential in cells of fibrous roots of sugar beet. Physiologia Plantarum 83: 54 – 62. Llugany M, Massot N, Wissemeier AH, Poschenrieder C, Horst WJ, Barcelo J. 1994. Aluminum tolerance of maize cultivars as assessed by callose production and root elongation. Journal of Soil Science and Plant Nutrition 157: 447–451. Ma QF, Rengel Z, Kuo J. 2002. Aluminium toxicity in rye (Secale cereale): root growth and dynamics of cytoplasmic Ca2+ in intact root tips. Annals of Botany 89: 241–244. Matsumoto H. 2000. Cell biology of aluminum toxicity and tolerance in higher plants. International Review of Cytology 200: 1– 46. Miyasaka SC, Kochian LV, Shaff JE, Foy CD. 1989. Mechanisms of aluminum tolerance in wheat. An investigation of genotypic differences in rhizosphere pH, K+, and H+ transport, and root-cell membrane potentials. Plant Physiology 91: 1188–1196. Montero M, Garcia-Sancho J, Alvarez J. 1994. Activation by chemotactic peptide of a receptor-operated Ca2+ entry pathway in differentiated HL60 cells. Journal of Biological Chemistry 269: 29451–29456. Monzer J. 1991. Ultra-structure of secondary plasmodesmata formation in regenerating Solanum nigrum protoplast cultures. Protoplasma 165: 86 – 95. Nakamura C, Telgen HV, Mennes AM, Ono H, Libbenga KR. 1988. Correlation between auxin resistance and the lack of a membrane-bound auxin binding protein and a root-specific peroxidase in Nicotiana tabacum. Plant Physiology 88: 845 – 849. Nichol BE, Oliveira LA. 1995. Effects of aluminium on the growth and distribution of calcium in roots of an aluminum-sensitive cultivar of barley (Hordeum vulgare). Canadian Journal of Botany 73: 1849–1858. Pouliquin P, Grouzis JP, Gibrat R. 1999. Electrophysiological study with Oxonol VI of passive NO3− transport by isolated plant root plasma membrane. Biophysical Journal 76: 360–373. Price AH, Tailor A, Ripley SJ, Griffiths A, Trewavas AJ, Knight MR. 1994. Oxidative signals in tobacco increase cytosolic calcium. Plant Cell 6: 1301–1310. Rengel Z. 1992a. Role of calcium in aluminium toxicity. New Phytologist 121: 499–513. Rengel Z. 1992b. Disturbance of Ca2+ homeostasis as a primary trigger in the aluminum toxicity syndrome. Plant, Cell & Environment 15: 931–938.


Rengel Z. 1996. Tansley review 89. Uptake of aluminium by plant cells. New Phytologist 134: 389–406. Rengel Z, Zhang WH. 2003. Tansley review. Role of dynamics of intracellular calcium in aluminium toxicity syndrome. New Phytologist 159: 295–314. Ryan PR, Kochian LV. 1993. Interaction between aluminum toxicity and calcium uptake at the root apex in near-isogenic lines of wheat (Triticum aestivum L.) differing in aluminum tolerance. Plant Physiology 102: 975–982. Siegel N, Haug A. 1983. Aluminum interaction with calmodulin. Evidence for altered structure and function from optical and enzymatic studies. Biochimica et Biophysica Acta 744: 35–45. Sivaguru M, Horst WJ. 1998. The distal part of the transition zone is the most aluminum-sensitive apical root zone of maize. Plant Physiology 116: 155–163. Sivaguru M, Baluska F, Volkmann D, Felle HH, Horst WJ. 1999a. Impacts of aluminum on the cytoskeleton of the maize root apex. Short-term effects on the distal part of the transition zone. Plant Physiology 119: 1073–1082. Sivaguru M, Yamamoto Y, Matsumoto H. 1999b. Differential impacts of aluminium on microtubule organization depends on growth phase in suspension-cultured tobacco cells. Physiologia Plantarum 10: 110–119. Sivaguru M, Fujiwara T, Samaj J, Baluska F, Yang Z, Osawa H, Maeda T, Mori T, Volkmann D, Matsumoto H. 2000a. Aluminum-induced 1,3-β--glucan inhibits cell-to-cell trafficking of molecules through plasmodesmata. A new mechanism of aluminum toxicity in plants. Plant Physiology 124: 991–1005. Sivaguru M, Matsumoto H, Horst WJ. 2000b. Control of responses to aluminum toxicity. In: Nick P, ed. Plant microtubules, potential use in plant biotechnology. Berlin, Germany: Springer Verlag, 103–120. Sivaguru M, Pike S, Gassmann W, Baskin T. 2003. Aluminum rapidly depolymerizes cortical microtubules and depolarizes the plasma membrane: evidence that these responses are mediated by a glutamate receptor. Plant Cell Physiology 44: 667–675. Tabuchi A, Matsumoto H. 2001. Changes in cell-wall properties of wheat (Triticum aestivum) roots during aluminum-induced growth inhibition. Physiologia Plantarum 112: 353–358. Taylor GJ, McDonald-Stephens JL, Hunter DB, Bertsch PM, Elmore D, Rengel Z, Reid RJ. 2000. Direct measurement of aluminum uptake and distribution in single cells of Chara corallina. Plant Physiology 123: 987–996. Teraoka T, Kaneko M, Mori S, Yoshimura E. 2002. Aluminum rapidly inhibits cellulose synthesis in roots of barley and wheat seedlings. Journal of Plant Physiology 159: 17–23. Vitorello VA, Haug A. 1996. Growing tobacco cells respond to rapid medium changes with a transient increase in localized Ca2+ activity. Plant Cell Reporter 16: 63–66. Vitorello VA, Haug A. 1997. An aluminum–morin fluorescence assay for the visualization and determination of aluminum in cultured cells of Nicotiana tabacum L. cv. BY-2. Plant Science 122: 35–42. Wissemeier AH, Horst WJ. 1995. Effect of calcium supply on aluminium-induced callose formation, its distribution and persistence in roots of soybean (Glycine max (L.) Merr.). Journal of Plant Physiology 145: 470–476. Wissemeier AH, Klotz F, Horst WJ. 1987. Aluminum induced callose synthesis in roots of soybean (Glycine max L.). Journal of Plant Physiology 129: 487–492. Yamamoto Y, Rikiishi S, Chang Y-C, Ono K, Kasai M, Matsumoto H. 1994. Quantitative estimation of aluminium toxicity in cultured tobacco cells: correlation between aluminium uptake and growth inhibition. Plant Cell Physiology 35: 575–583. Yamamoto Y, Hachiya A, Matsumoto H. 1997. An aluminum-tolerant cell line isolated from cultured tobacco cells. In: Ando T, Fujita K, Mae T, Matsumoto H, Mori S, Sekiya J, eds. Plant nutrition – for sustainable


Research food production and environment, Dordrecht, The Netherlands: Kluwer Academic Publishers, 451– 452. Yermiyahu U, Brauer DK, Kinraide TB. 1997. Sorption of aluminum to plasma membrane vesicles isolated from roots of Scout 66 and Atlas 66 cultivars of wheat. Plant Physiology 115: 1119 –1125. Zhang G, Hoddinott J, Taylor GJ. 1994. Characterization of 1,3-β--glucan (callose) synthesis in roots of Triticum aestivum in response to aluminum toxicity. Journal of Plant Physiology 144: 229–234.

Zhang WH, Rengel Z. 1999. Aluminium induces an increase in cytoplasmic calcium in intact wheat root apical cells. Australian Journal of Plant Physiology 26: 401–409. Zhang WH, Rengel Z, Kuo J. 1998. Determination of intracellular Ca2+ in cells of intact wheat roots: loading of acetoxymethyl ester of Fluo-3 under low temperature. Plant Journal 15: 147–151. Zhang WH, Rengel Z, Kuo J, Yan G. 1999. Aluminium effects on pollen germination and tube growth of Chamelaucium uncinatum. A comparison with other Ca2+ antagonists. Annals of Botany 84: 559–564.

About New Phytologist • New Phytologist is owned by a non-profit-making charitable trust dedicated to the promotion of plant science, facilitating projects from symposia to open access for our Tansley reviews. Complete information is available at www.newphytologist.org. • Regular papers, Letters, Research reviews, Rapid reports and Methods papers are encouraged. We are committed to rapid processing, from online submission through to publication ‘as-ready’ via OnlineEarly – the 2003 average submission to decision time was just 35 days. Online-only colour is free, and essential print colour costs will be met if necessary. We also provide 25 offprints as well as a PDF for each article. • For online summaries and ToC alerts, go to the website and click on ‘Journal online’. You can take out a personal subscription to the journal for a fraction of the institutional price. Rates start at £109 in Europe/$202 in the USA & Canada for the online edition (click on ‘Subscribe’ at the website). • If you have any questions, do get in touch with Central Office ([email protected]; tel +44 1524 592918) or, for a local contact in North America, the USA Office ([email protected]; tel 865 576 5261).



109