Journal of Experimental Botany, Vol. 53, No. 376, pp. 1959-1966,
September 1, 2002
© 2002 Oxford University Press
Aluminium-induced growth inhibition is associated with impaired efflux and influx of H+ across the plasma membrane in root apices of squash (Cucurbita pepo)
Received 3 January 2002; Accepted 5 June 2002
1 Research Institute for Bioresources, Okayama University, Chuo 2-20-1, Kurashiki 710-0046, Japan
2 Department of Horticulture, Agricultural Plant Stress Research Center, College of Agriculture, Chonnam National University, Kwangju 500-757, Korea
3 Soil Science and Plant Nutrition, The University of Western Australia, 35 Stirling Highway, Crawley WA 6009, Australia
4 To whom correspondence should be addressed: Fax: +81 86 434 1249. E-mail: hmatsumo{at}rib.okayama-u.ac.jp
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Abstract |
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It is generally understood that the inhibition of growth of root apices is the initial effect caused by aluminium (Al) toxicity. The correlation between impaired H+-fluxes across the plasma membrane (PM) and Al-induced growth inhibition, Al accumulation and callose formation in root apices of squash (Cucurbita pepo L. cv. Tetsukabuto) is reported here. The root inhibition was dependent on Al concentration, and the duration of exposure, with the damage occurring preferentially in regions with high Al accumulation and callose formation. Using the fluorescent Al indicator (Morin), Al was localized in the cell walls of the root-tip cells after 3 h and in the whole root-tip cells after 6 h of the Al treatment (50 µM). The inhibition of H+-pumping rate in the highly purified PM vesicles obtained from the Al-treated apical root portions (1 cm) coincided with the inhibition of root growth under Al stress. Furthermore, H+-ATPase activity of PM vesicles prepared from the control root apices was strongly inhibited by Al in vitro in a dose-dependent manner. Approximately 50% inhibition was observed when PM vesicles were preincubated at Al concentration as low as 10 µM followed by the enzyme assay in the medium without Al. Using the pH indicator (bromocresol purple), it is shown that surface pH of the control (0 Al) root apices was strongly alkalized from the starting pH of 4.5 in a time-dependent manner. By contrast, the surface pH changed only slightly in the Al-treated root apices. The changes in surface pH mediated by altered dynamics of H+ efflux and influx across the root tip PM play an important role in root growth as affected by Al.
Key words: Key words: Al accumulation, Al toxicity, callose formation, Cucurbita pepo, H+ efflux, H+ influx, plasma membrane, surface pH.
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Introduction |
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It has been well established that aluminium (Al) inhibits root growth rapidly, with the root apex being the primary target playing a central role in Al toxicity and tolerance (Horst, 1995; Kochian, 1995; Matsumoto, 2000; Taylor, 1995, for recent reviews). However, the mechanism of Al toxicity in plants still remains poorly understood.
Until now, several hypotheses have been suggested for Al toxicity mechanisms, such as changes in cytosolic free Ca2+ (Jones et al., 1998; Zhang and Rengel, 1999), rapid callose formation (Horst et al., 1997), and alterations in root cytoskeleton (Sivaguru et al., 1999). In addition, several studies have focused on the plasma membrane (PM) functions, while the potential apoplastic and symplastic Al target sites in plant cells are under debate (Horst, 1995; Kochian, 1995; Rengel, 1996). PM is an important barrier for the passive movement of Al. Some of the PM properties, such as surface negativity, are altered rapidly; these changes have frequently been suggested to be the basis for the observed differences in Al tolerance/toxicity between plant genotypes (Ahn et al., 2001; Kinraide, 1994; Kinraide et al., 1998; Miyasaka et al., 1989; Wagatsuma and Akiba, 1989; Yermiyahu et al., 1997).
At the cellular level, proton influx occurs at the root apical (meristematic) zone, whereas the elongation zone exhibits proton efflux (Piñeros and Kochian, 2002). The H+-pumping (H+-ATPase activity) across the PM is supposed to play a major role in cytoplasmic pH regulation. The H+-ATPase activity is strongly pH dependent, with an optimal pH around 6.6. As a consequence, a marked stimulation of H+-ATPase occurs when cytoplasmic pH is lowered. Also, in many cases enhanced H+-pumping activity results in apoplastic acidification (Morsomme and Boutry, 2000).
An important property of plants is their ability to change the pH of the rhizosphere during growth. Many reports have tried to correlate Al resistance and transient increases in the pH of the rooting medium. For example, Foy et al. (1965) proposed an Al-exclusion mechanism due to an increase in rhizosphere pH. Alkalization of the rhizosphere may reduce the concentration of Al3+ in favour of less-toxic Al species such as Al hydroxide, because the solubility of Al is strongly dependent on pH. In most of these experiments, pH measurement was done in bulk solution, which did not permit accurate assessment of pH at the root surface. Recently, Degenhardt et al. (1998) and Kollmeier et al. (2000) found a clear Al-induced increase in pH by using a vibrating pH-sensitive microelectrode at the root tip surface. Other in vivo techniques (e.g. using agar supplemented with specific reagents and indicators) have been developed for demonstration of chemical changes along roots in nutrient solution or in soil. Calba et al. (1996) demonstrated that agarose is a more suitable medium than agar for studying rhizosphere Al dynamics, because agarose is nearly devoid of phosphorus and other Al-complexing substances.
In this study, an investigation was undertaken of the interrelationship between growth inhibition, accumulation of Al and callose, and changes of H+ efflux and H+ influx in squash root apices under Al stress.
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Materials and methods |
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Growth conditions and root growth measurements
Squash seeds (Cucurbita pepo L. cv. Tetsukabuto) were soaked in tap water for 12 h and germinated in an incubator (28 °C) in the dark for 24 h on two layers of filter paper saturated with one-fifth Hoagland solution (HS) adjusted to pH 4.5 with 0.1 M HCl. After germination, uniform seedlings with 1.5 cm root length were transferred to polyethylene pots and grown in a controlled-environment chamber with a 14/10 h day/night regime under 580 µmol m–2 s–1 of light during the day. The day/night temperatures were set at 25/20 °C with 65% constant relative humidity. The HS was continuously aerated and was replaced every day to minimize pH change. Five-day-old seedlings were transferred to a modified HS without phosphate at least 12 h prior to Al treatment of 0, 20, 50 or 100 µM AlCl3.6H2O (pH 4.5). The primary root length of 10 seedlings was measured with a ruler at 0, 3, 6, and 24 h after the commencement of Al treatment. In a parallel experiment, seedlings were washed in double-distilled water (DDW) for 5 min, stained with a 0.1% (w/v) aqueous solution of Eriochrome cyanine R (Sigma, Tokyo, Japan) for 10 min, washed in DDW and photographed using a light microscope. Eriochrome cyanine R stains Al (Aniol, 1983).
Al localization and callose contents in root apex
After the Al treatments mentioned above, the roots were washed in DDW, and 1 cm apical portions (including the root cap) were cut using a razor blade. The excised root apices (four replicate segments) were transferred to 1.5 ml Eppendorf (Netheler-Hinz Gmbh, Hamburg, Germany) tubes each containing 1 ml of 2 M HCl for 48 h. The Al content in the HCl digests was determined by an atomic absorption spectrophotometer (Z-8270, Hitachi, Tokyo, Japan) after dilution.
Seedlings from the parallel experiments were washed for 5 min in DDW, stained with 10 mM MES [2-(N-morpholino)ethanesulphonic acid] buffer (pH 5.5) containing 100 µM Morin (Sigma, Tokyo, Japan) for 30 min. After a further wash in MES buffer, the images were obtained using a Zeiss confocal microscope (Axioplan 2 connected with LSM 510, Carl Zeiss, Oberkochen, Germany) at 488 nm (Argon laser) excitation wavelength.
For quantifying callose, apical root portions (2 cm) were fixed in 96% ethanol, briefly washed and cut into 1 cm pieces starting from the tip. After dissection, the segments were blotted dry and transferred immediately to Eppendorf tubes containing 1 M NaOH. Callose levels were estimated following the Kauss (1996) method. Briefly, each sample containing similarly treated root segments in NaOH was homogenized directly for 1 min. Subsequently, the samples were placed in a water bath (80 °C, 30 min) to solubilize the callose and centrifuged (15 min, 12 000 g) at room temperature. Callose concentration in the supernatant was quantified fluorimetrically at 393 nm excitation and 484 nm emission wavelengths (Hitachi-4500 fluorescence sprectrophotometer, Tokyo, Japan) with the decolorized aniline blue technique using Curdlan as reference.
Preparation of PM vesicles
Plasma membrane (PM) vesicles were prepared strictly at 4 °C following the method of Palmgren et al. (1990). Briefly, after Al treatment, the 1 cm apical segments from primary roots (approximately 4 g fresh weight) were ground in the presence of insoluble polyvinylpyrrolidone with a homogenizing buffer containing 330 mM sucrose, 50 mM MOPS-1,3-bis(tris[hydroxymethyl]methylamino) propane (BTP), pH 7.5, 5 mM EDTA, 5 mM dithiothreitol (DTT), 0.5 mM phenylmethysulphonyl fluoride, 0.2% (w/v) bovine serum albumin (BSA) (Sigma, Tokyo, Japan, protease free), and 0.2% (w/v) casein. The homogenate was filtered through four layers of cheesecloth and centrifuged (10 000 g, 20 min). The supernatant was ultra-centifuged (100 000 g, 1 h) and the resulting precipitate was suspended with a glass homogenizer in suspension buffer consisting of 330 mM sucrose, 5 mM K-phosphate (pH 7.8), 5 mM KCl, 1 mM DTT, and 0.1 mM EDTA. The homogenate was loaded on a 12 g two-phase system, containing 6.5% (w/w) Dextran T500, 6.5% (w/w) polyethylene glycol 3350, 330 mM sucrose, 5 mM K-phosphate (pH 7.8), 5 mM KCl, 1 mM DTT, and 0.1 mM EDTA. After the batch procedure, the resulting upper phase was mixed with a dilution buffer that consisted of 330 mM sucrose, 5 mM MOPS-BTP (pH 7.5), 5 mM KCl and centrifuged (100 000 g, 1 h). The PM vesicles were used immediately or stored at –80 °C until further analysis.
Determination of PM H+-pumping
Proton uptake into the inside-out vesicles (5 µg of protein) was monitored as the decrease in the intensity of the acridine orange at 495 nm. The assay medium consisted of 20 µM acridine orange, 2 mM ATP–BTP (pH 7.0), 140 mM KCl, 4 mM MgCl2, 1 mM EDTA, 1 mM DTT, 1 mg ml–1 BSA (essentially fatty acid free), and 50 µg PM protein in a total volume of 2 ml. Fifty µM gramicidine was added as indicated. After 5 min preincubation at 20 °C, the reaction was initiated by addition of ATP. The rate of H+ accumulation was estimated from the initial slope of absorbance quenching (
A495) of acridine orange.
Determination of PM H+-ATPase activity after in vitro Al treatment
The highly purified right-side-out PM vesicles (5 µg) were treated with 0, 1, 2, 5, 10, 50, and 100 µM Al in vitro for 10 min and centrifuged (100 000 g, 1 h) to remove the unbound Al. H+-ATPase activity was measured in an assay system containing 50 mM MOPS–BTP (pH 6.5), 2.5 mM MgSO4, 50 mM KCl, 2.5 mM TRIS–ATP, and 0.05% (w/w) Brij 58 (polyoxyethylene 20 cetyl ether, Sigma, Tokyo, Japan) to produce inside-out vesicles (Johansson et al., 1995) and an appropriate amount of H+-ATPase. The reaction was carried out for 30 min at 37 °C. Five hundred microlitres of 5% (w/v) cold TCA and 2 ml of 0.1 M Na-acetate was added to the mixture and centrifuged (2000 g, 10 min) after the addition of 0.3 ml of 1% (w/v) ammonium molybdate in 0.025 M H2SO4. The Pi liberated during 10 min at 30 °C was measured with a spectrophotometer (Model UV-1201; Shimadzu, Tokyo, Japan) at 720 nm. The membrane protein was determined with the Bradford (1976) method using BSA as standard.
Visualizing surface pH
To visualize alkalinization along single intact roots in the presence or absence of Al in situ, a modified agarose plate technique was used instead of agar as the culture medium because agarose is a relatively pure substrate for studying rhizosphere Al dynamics (Calba et al., 1996). Low-gelling agarose (0.7%, w/v), dissolved in one-fifth HS without phosphate, was poured into a 150 ml vertical plate (diameter 20 cm) containing pH indicator Bromocresol Purple (0.015%, w/v). The solution was adjusted to pH 4.5 before being poured into the plate, and five roots were embedded into agarose just before solidification. For the Al treatments, 180 µM Al was added to the agarose solution to achieve a final monomeric Al concentration of 50 µM according to the Kerven et al. (1989) method. The pH values were obtained by comparing a standard coloured scale in the same plate, but without plants (range of colour pH scale: 4.0–6.5 with intervals of 0.5 pH unit). Each experiment was carried at least three times with four replicates.
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Pattern of root growth inhibition by Al
The plants were grown in one-fifth HS adjusted to pH 4.5 for 5 d after germination. Al treatments were performed in the same solution (but without P); the plants were cultured in the –P solution for at least 12 h prior to treatment. As a first approach, the growth inhibition of primary roots was measured. Within the initial 3 h of Al treatment, significant growth inhibition occurred only at 100 µM Al treatment. Roots treated with 10 and 20 µM Al were not inhibited until 6 h. In the 50 µM Al treatment, a significant inhibition (45%) of root growth was observed at 6 h (Fig. 1). Eriochrome cyanine R staining was markedly observed only in the elongation zone (the region approximately 2–4 mm from the root apex) in the treatments with 50 and 100 µM Al at 6 h (Fig. 2). Roots treated with high Al concentrations (50 and 100 µM) for 24 h were severely affected, becoming thick, stubby and dark compared with control roots in the root tip region (data not shown). Therefore, all subsequent experiments were performed with roots grown in the presence or absence of 50 µM Al.
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Localization and quantification of Al and Al-induced callose
Localization of Al using an Al-specific stain, Morin, coupled with confocal laser scanning microscopy revealed that most of the Al was localized preferentially in the cell walls after 3 h of Al treatment (Fig. 3, set A). However, after 6 h, Al were detected in the whole cells, especially in the apex (Fig. 3B") and the basal root regions that were severely damaged (Fig. 3B). Furthermore, the 2–4 mm region showed the most severe growth inhibition in response to Al. In this region Al accumulation was higher in the cortex than the epidermal cells (Fig. 3C, D). In order to compare the Al-staining pattern and the Al-induced growth inhibition, Al contents from the root apex (1 cm) were quantified after Al treatment. High Al contents coincided with intensive staining for Al and reduced growth after Al treatments (data not shown). A possibility cannot be excluded that Al accumulation occurred in cells that died from Al damage.
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Al-induced callose levels were highest in root segments 4–6 mm from the root apex after the 3 h Al treatment. However, after 6 h, the segment 6–8 mm from the root tip showed the peak in callose level, indicating that the Al-induced damage in the segment 4–6 mm from the root apex moved backwards after 6 h, which corresponds to approximately 2 mm growth after this time (Fig. 4). Although the longer duration of Al treatment induced a higher amount of callose, the magnitude of callose differences between the 3 h and 6 h treatments was not large (data not shown).
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Effect of Al on PM H+-pumping and H+-ATPase activity
The purity of PM vesicles was first validated using vesicles collected from the 1 cm root apices in the presence or absence of various inhibitors. Vanadate, which is a specific inhibitor of PM ATPase activity, inhibited about 90% of the activity, while nitrate and azide resulted in no inhibition. The orientation of PM vesicles was determined by measuring ATPase latency. A huge increase in H+-ATPase activity was observed with the addition of Brij 58, indicating that the vesicles were sealed, right-side-out and of high purity (data not shown).
Since the region about 1–4 mm from the root tip was the most sensitive site to the treatment by 50 µM Al after 6 h (Fig. 2), roots were treated with 50 µM Al concentration for 3, 6, and 24 h and right-side-out PM vesicles from the 1 cm apical root portions were isolated. H+-pumping activity was measured in the presence of 0.05% (w/v) Brij 58. Al severely inhibited the H+-transport rates by inside-out root tip PM vesicles isolated after 3 h (20%) and 6 h (55%) as compared to respective control root tips (Fig. 5). Gramicidine at 50 µM dissipated the pH, and the quenching of acridine orange did not occur without ATP.
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The in vitro effect of Al on the H+-ATPase activity was analysed using highly purified right-side-out PM vesicles from the control root tips. The PM vesicles (5 µg) in a dilution buffer (pH 7.5) were treated with 0, 1, 2, 5, 10, 50 or 100 µM Al in vitro for 10 min and centrifuged to remove the unbound Al. The inhibition of the H+-ATPase activity was dose-dependent and was significant even at 1 µM. Approximately 50% inhibition occurred at concentration as low as 10 µM Al, and complete inhibition at 50 and 100 µM (Fig. 6).
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Changing patterns of surface pH by Al
Colour changes due to surface pH were monitored using a modified agarose plate technique and the pH indicator bromocresol purple in intact roots treated with 50 µM Al for 3, 6 or 24 h. The patterns of surface pH were not only visualized, but also quantified by comparing a standard colour scale (see Materials and methods) after 3, 6 and 24 h in the presence or absence of 180 µM Al [to achieve a final monomeric Al concentration of 50 µM according to the Kerven et al. (1989) method] at pH 4.5. Alkalization along the root surface was detected by observing colour change from yellow to purple (Fig. 7). The change in surface pH was detectable after 3 h in the control root. The strong pH increase (from pH 4.5 to 6.0) was visible in the 1 cm root zone after 24 h. In Al-treated roots, alkalization was detectable just behind the root tip only after 24 h (Fig. 7B"). The pH of 2.0 l bulk solution (one-fifth HS concentration, starting pH 4.5) containing five plants was also determined. The changes of pH in bulk solution were negligible after 6 h, but after 24 h, pH values of control and Al-treated solution were 5.1 and 4.7, respectively (data not shown).
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Discussion |
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Al-induced growth inhibition in the apex
Aluminium (50 µM) inhibited root apical growth within 3 h and the inhibition was significant at 6 h (Figs 1, 2). The pattern of Al inhibition observed in this study was similar to the inhibitory effect of Al reported for squash roots under the high Al concentrations (1 mM) in the nutrient solution ((Le Van et al., 1994). In the case of corn, the inhibition of total root elongation matched the pattern of segmental inhibition (Blancaflor et al., 1998). An earlier spatial analysis of root growth suggested that the 2–4 mm is the central elongation zone for squash (Ahn et al., 2001).
The results of Morin staining indicated that the Al accumulation was greater in the cortex than in epidermal cells of the same region (Fig. 3), which raises the question of the use of surface stains to study the pattern of Al accumulation. After 6 h Al treatment, there was an increase in damage in the root apical surface cells, resulting in enhanced uptake of Al by cells. This led to severe physical damage to the more mature regions of the root apex, which indicates that there may be spatial differences in Al sensitivity of cells in the root apex. Ryan et al. (1993) described similar results, showing higher Al injury to basal zone cells of corn roots.
Al-induced callose formation has often been associated with the inhibition of root growth in several monocot plants, including wheat and corn (see Horst, 1995, for a review). For instance, increased Al accumulation caused growth inhibition and callose formation in a narrow zone (apical 2 mm segment) of wheat (Rincon and Gonzales, 1992; Samuels et al., 1997; Tice et al., 1992) and in a 1 mm segment of corn (Sivaguru and Horst, 1998). The present study with squash, however, suggests that the maximum Al-induced callose accumulation does not coincide with the pattern of Al-induced growth inhibition and Al accumulation (Fig. 4). The present results may reflect the intrinsic differences between monocot and dicot plants in the cell wall composition (such as pectin and xyloglucans), content and architecture (Carpita and Gibeaut, 1993) and their differential response to Al.
Al-related impairment of H+ efflux and H+ influx across PM
Micromolar concentrations of Al can effectively interfere with the H+-ATPase activity of PM-enriched vesicles, which is related to trans-membrane potential (Hamilton et al., 2001; Matsumoto, 1988; Matsumoto et al., 1992; Sasaki et al., 1994; Siegel and Haug, 1983). Recently, Ahn et al. (2001) reported that this mechanism is related to the inhibition of H+ efflux through the PM due to depolarization of surface potential under Al stress. In agreement with these findings, it is reported here that a close association exists between Al-induced root growth inhibition and decreased H+-pumping activity under Al toxicity in vivo (Fig. 5) and in vitro (Fig. 6). The activity of this enzyme plays a central role in the functional association of PM surface charge and H+ efflux, and is a crucial factor for the survival of plants under various environmental stresses. For instance, a decrease in PM surface potential (depolarization) is correlated with the decline of H+-ATPase activity in PM vesicles of plant roots under low temperature (Ahn et al., 2000), salt stress (Suhayda et al., 1990), and Al toxicity (Ahn et al., 2001). In addition, using bromocresol purple in agarose gel, a more vigorous H+ influx into root apices of the control than Al-treated roots (Fig. 7) was found. This is in full agreement with (i) the decreased H+ efflux from the root tips of Al-sensitive wheat (Miyasaka et al., 1989), and (ii) an increased H+ influx into the root apices with enhanced Al tolerance in the mutant plants of Arabidopsis subjected to Al (Degenhardt et al., 1998).
Al3+ has a very strong affinity for the PM surface (e.g. 56-fold higher than for Ca2+) (Akeson et al., 1989). Ahn et al. (2001) reported that Al at 50 µM could neutralize the surface charge of the PM and cause a surface potential shift from –20 to +1 mV. Such an Al-related shift in PM surface potential causes disturbance in ion transport processes. In an Al-sensitive cultivar of barley, 100 µM Al inhibited the influx of Ca2+ (69%), NH4+ (40%), and K+ (13%) and enhanced the influx of NO3– (44%) and phosphate (17%) (Nichol et al., 1993). Therefore, a possible mechanism is proposed whereby Al binds to the PM at the root tips, shifting the surface potential to more positive values and thus influencing ion movement at the binding sites of transport proteins. A positively charged layer at the outside PM surface of Al-treated root tips would retard the uptake of cations, including H+ (as documented in this study), causing less alkalization around the root tips compared with the control roots not treated with Al. It should be borne in mind that H+ influx at the root tip, and thus alkalization, represents normal root functioning under physiological conditions (Piñeros and Kochian, 2002). However, the Al-related inhibition of the H+-ATPase activity and hence decreased pumping of H+ out of the cell, as well as an expected increased uptake of anions by the Al-treated root tips and thus likely increased H+ influx via co-transport, would tend to increase alkalization of the medium around Al-treated root tips, thus arguing against the proposed hypothesis. Apparently, the magnitude of the ion fluxes mentioned would need to be determined first in order to understand the effects that individual fluxes may have on apparent alkalization of the environment around the root tips. Further research on determining simultaneously the H+ influx and the H+ efflux, as well as cation and anion fluxes, at the root tips is required to explain the role of H+ fluxes in Al toxicity and tolerance fully.
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Acknowledgements |
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This study was supported by the Program for the Promotion of Basic Research Activities in Innovative Biosciences (PROBRAIN) to HM and a Grant-in-Aid for general research, Ministry of Agriculture, Forests and Fisheries, Japan. Scientific Research (A) (No. 09460038 to HM) from the Ministry of Education, Science, Sports and Culture of Japan, and the Ohara Foundation for Agricultural Science. We thank Professor Yoko Yamamoto (Research Institute for Bioresources, Kurashiki, Japan) and Professor Walter Horst (Hannover University, Germany) for fruitful discussions at the start of this study and Ms Michiyo Ariyoshi for her kind help during the course of the experiments.
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