JXB Advance Access originally published online on June 1, 2007
Journal of Experimental Botany 2007 58(10):2565-2572; doi:10.1093/jxb/erm107
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RESEARCH PAPER |
Microtubule dynamics in relation to osmotic stress-induced ABA accumulation in Zea mays roots
1College of Bioscience and Biotechnology, Key Laboratory of Crop Genetics and Physiology of Jiangsu Province, Yangzhou University, Yangzhou, 225009 People's Republic of China
2Department of Biology, Hong Kong Baptist University, Hong Kong, China
* To whom correspondence should be addressed. E-mail: jsliang{at}mail.yzu.edu.cn
Received 15 February 2007; Revised 31 March 2007 Accepted 4 April 2007
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Abstract |
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Microtubules play important roles in many physiological processes such as plant responses to drought stress. Abscisic acid (ABA) accumulates significantly in plants in response to drought conditions, which has been considered as a major response for plants to enhance drought tolerance. In this work, the focus was on the possible roles of microtubules in the induction of ABA biosynthesis in the roots of Zea mays when subjected to osmotic stress. The dynamic changes of microtubules in response to the stress were investigated by immunofluorescence staining, enzyme-linked immunosorbent assay, and a pharmacological approach. Disruption and stabilization of microtubules both significantly stimulated ABA accumulation in maize root cells, although this stimulation was markedly lower than that caused by osmotic stress. Cells in which the microtubule stability had been changed did not respond further to osmotic stress in terms of ABA biosynthesis. However, treatment with both a microtubule de-stabilizer and a stabilizer enhanced the sensitivity of cells to osmotic stress in terms of ABA accumulation. It is suggested that both osmotic stress and changes in microtubule dynamics would trigger maize root cells to biosynthesize ABA, and interactions between osmotic stress and microtubule dynamics would have an effect on ABA accumulation in root cells, although the exact mechanism is not clear at present.
Key words: ABA biosynthesis, microtubules, osmotic stress, Zea mays
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Introduction |
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Microtubules are cylindrical polymers comprised mainly of
- and β-tubulin heterodimers. An individual microtubule is about 24 nm in diameter and up to 10 µm in length. However, many microtubules may overlap, cross-bridge, or interact with one another to form large and highly ordered arrays (Baskin and Cande, 1990). In higher plants, at least four distinct arrays have been identified, i.e. the interphase cortical array, the pre-prophase band, the mitotic spindle, and the phragmoplast (Goddard et al., 1994). Among them, cortical microtubules are essential for normal plant morphogenesis because they determine the placement of cell division planes and affect the axes of cell elongation. During interphase, a cortical array consisting of parallel microtubules oriented perpendicularly to the cell expansion axis assists cellulose deposition (Cyr, 1994) and responds to stimuli (Wymer et al., 1996). As an important component of cellular morphogenesis, plant microtubules have been shown to be essential for embryo development (Steinborn et al., 2002), organ formation (Whittington et al., 2001), organ twisting (Thitamadee et al., 2002), and stomatal movement (Marcus et al., 2001). Furthermore, the function of the cortical microtubules is intimately linked to their organizational state, which is subject to spatial and temporal modifications by developmental and environmental cues (Dixit and Cyr, 2004). Drought stress, one of the main adverse environmental factors, regulates many aspects of plant growth and development. Many adaptive strategies have evolved in plants for coping with drought stress, among which abscisic acid (ABA) accumulation in plant cells upon water deficit is one of the most important ways of enhancing plant drought tolerance. ABA can induce stomatal closure and stimulate water uptake by roots, and thus maintain water balance in the plant body. Water stress readily leads to plant tissue dehydration, characterized by a decreased turgor pressure and cell volume (Boudsocq and Laurière, 2005). Dehydration triggers plasmolysis of cells and accelerated biosynthesis of ABA in both leaves and roots (Jia et al., 2001). Plasmolysis also destroys the microtubule cytoskeleton and the geometry of the wall, and thus could provide spatial information to which the microtubule cytoskeleton responds (Pollock and Pickett-Heaps, 2005). Destruction of microtubules by hyperosmotic stress and their replacement by tubulin microtubules and putative tubulin paracrystals are common features among angiosperms. Thus, microtubules were suggested to be involved in the mechanism of protoplast volume regulation (Komis et al., 2002).
It was demonstrated that ABA produced steeply oblique microtubule bundles in Chinese winter wheat (Triticum aestivum L.) (Wang and Nick, 2001). Also, ABA disrupted cortical microtubules in guard cells, but not in epidermal cells, with concomitant closure of stomata. Thus, it was suggested that disruption of microtubules seems to be a specific effect of ABA in guard cells, but its physiological significance remains obscure (Jiang et al., 1996). In the last decade, plenty of information has become available about the linkage between microtubules and ABA in plant cells, but the mechanisms involved and the responses of plant microtubules to drought stress remain largely unknown.
Usually dynamic microtubules are more sensitive than stable microtubules to extracellular stimuli (Marcus et al., 2001), and drought stress affects both the microtubule dynamics and ABA accumulation. Therefore, it is reasonable to assume that microtubule dynamics may be involved in the stimulation of ABA biosynthesis under drought stress conditions. In the present study, this hypothesis was tested by using microtubule-destabilizing (e.g. oryzalin) and -stabilizing (e.g. taxol) reagents, combined with osmotic stress treatment, to investigate if microtubule dynamics regulate ABA biosynthesis in maize roots under osmotic stress. The results indicated that microtubule dynamics and ABA accumulation are significantly affected by osmotic stress, and changes of microtubule dynamics caused by microtubule-destabilizing and -stabilizing reagent treatments resulted in a significant change in ABA accumulation. It was suggested that the stimulation of ABA biosynthesis under an osmotic stress condition may be through changes in microtubule dynamics. The possible mechanism of changes in ABA accumulation caused by microtubule dynamics is also discussed.
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Materials and methods |
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Plant materials and growth conditions
Maize (Zea mays cv. Xiyu 3) seeds were germinated under controlled conditions in a growth chamber at a temperature of 28 °C for 5 d. The seedlings were then grown in a half-strength Hoagland culture solution under a 16 h light period supplied by cool-white fluorescent bulbs (100 µmol m–2 s–1) at a temperature of 30 °C and an 8 h dark cycle at 25 °C. Root segments of seedlings at the five-leaf stage were harvested for analyses.
Treatments of maize seedlings
Ten treatments of maize seedlings were conducted (Table 1), i.e. normal maize seedlings (as control), seedlings treated with 15% (w/v) polyethyleneglycol 6000 (PEG-6000) for different times, seedlings treated with 50 µmol l–1 oryzalin for 1 h, seedlings treated with 10 µmol l–1 taxol/paclitaxel for 24 h, seedlings treated with 50 µmol l–1 oryzalin for 1 h or 10 µmol l–1 taxol/paclitaxel for 24 h after being treated with 15% PEG for 12 h, and seedlings treated with 15% PEG for 12 h after being treated with 50 µmol l–1 oryzalin for 1 h or 10 µmol l–1 taxol/paclitaxel for 24 h.
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Immunofluorescence localization of
-tubulin in maize root cellsPrimary root segments (
1 cm from the tip) of maize seedlings at the five-leaf stage were harvested and fixed in 4% paraformaldehyde in phosphate-buffered saline (PBS, pH 7.0) at room temperature for 1 h and washed three times with PBS. The fixed root segments were enzymatically digested in PBS buffer containing 1% (w/v) cellulase and 0.5% (w/v) pectinase at 37 °C for 30 min. Cells harvested by centrifugation were treated with Triton X-100, then incubated in PBS buffer containing 1% (w/v) bovine serum albumin (BSA) for 10 min. Cells were then incubated with a primary antibody against mouse -tubulin (T5168, Sigma, St Louis, MO, USA) at a dilution of 1:2000 at 37 °C for 2 h. After washing three times with PBS, cells were incubated with fluorescein isothiocyanate (FITC)-conjugated anti-mouse immunoglobulin (whole-molecule; F0257, Sigma) at a dilution of 1:400 at 37 °C for 2 h, then washed four times with PBS. Specimens were mounted in 2% DABCO (1,4-diazabicyclo[2,2,2,]octane; D-2522, Sigma) and examined using an Olympus BX 51 fluorescence microscope. FITC was excited with a 488 nm laser line, and FITC emission was detected using a 505–530 nm band-pass filter. A 100x oil-immersion objective (numerical aperture of 1.3) was used to scan the samples.
ABA extraction and quantification by enzyme-linked immunosorbent assay (ELISA)
The extraction and quantification of ABA were based on the procedure described by Chen et al. (2006). Briefly, the roots of maize seedlings prepared and treated as above were lyophilized and ground in an ice-cold mortar in 4 ml of 80% (v/v) methanol extraction medium containing 1 mmol l–1 butylated hydroxytoluene as an antioxidant. The extract was incubated at 4 °C for 24 h and centrifuged at 3000 g for 15 min at the same temperature. The supernatant was passed through Chromosep C18 columns (C18 Sep-Park Cartridge, Waters Corp., Millford, MA, USA), prewashed with 10 ml of 100% (w/v) and 5 ml of 80% (v/v) methanol, respectively. A 2 ml aliquot of hormone fractions eluted from the columns was dried under N2, and dissolved in 0.5 ml of PBS containing 0.1% (v/v) Tween-20 and 0.1% (w/v) gelatine (pH 7.5) for ABA analysis by competitive ELISA.
The antigens (ABA hapten-carrier protein), mouse monoclonal antibodies against ABA, and IgG horseradish peroxidase used in the ELISA were purchased from the Phytohormones Research Institute, China Agricultural University. ELISA was performed on a 96-well microplate. Each well on the plate was coated with 100 µl of coating buffer (1.5 g l–1 Na2CO3, 2.93 g l–1 NaHCO3, and 0.02 g l–1 NaN3, pH 9.6) containing 0.25 µg ml–1 antigens. The coated plates were incubated for 30 min at 37 °C, and then kept at room temperature for 3–4 min. After washing three times with PBS-Tween-20 [0.1% (v/v)] buffer (pH 7.4), each well was filled with 50 µl of either extracts or ABA standards (0–2000 ng ml–1 dilution range), and 50 µl of 20 µg ml–1 ABA antibodies. The plate was incubated for 30 min at 37 °C, and then washed as above. A 100 µl aliquot of 1.25 µg ml–1 IgG–horseradish peroxidase substrates was added to each well and incubated for 30 min at 37 °C. The plate was rinsed four times with the above PBS-Tween-20 buffer, and 100 µl of colour-producing solution containing 1.5 mg ml–1 o-phenylenediamine and 0.008% (v/v) H2O2 was added to each well. The reaction was stopped by adding 50 µl of 4 M H2SO4 per well when the 2000 ng ml–1 standard had a pale colour and the 0 ng ml–1 standard had a deep colour in the wells. Colour development in each well was detected using a Microplate Reader (model EL310, Bio-TEK, Winooski, VT, USA) at optical density A490. The results are the means ±SE of at least three replicates.
Quantitation of Vp14 gene expression by reverse transcription-PCR (RT-PCR)
Total RNA of maize roots prepared and treated as above was isolated using the RNeasy® Plant Mini Kit (QIAGEN). First strand cDNA was synthesized in a 25 µl reaction solution containing approximately 2 µg of total RNA using SuperScriptTM II Reverse Transcriptase (Invitrogen) and oligo(dT)18 as a primer. The synthesized first strand cDNA was used as a template. The primer pairs of Vp14 (GenBank accession no. ZMU95953) and actin are as follows: Vp14 forward primer 5’ TCCACGACTTCGCCATCACC 3', reverse primer 5’ CGTCTTCTCCTTGTCCAGCACC 3’; actin forward primer 5’ TGGGATGACATGGAGAAGAT 3', reverse primer 5’ ATACCAATCATAGATGGCTGG 3'.
PCR with 30 cycles was performed. The PCR products were subjected to 1% (w/v) agarose gel electrophoresis and stained with ethidium bromide.
Statistical analysis
The significance analyses were carried out using the SAS statistical analysis package (version 6.12, SAS Institute, Cary, NC, USA). Multiple comparisons were made among different treatments.
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Results |
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Impacts of hyperosmotic stress (PEG-6000 treatment) on
-tubulin arrangement and ABA content of maize root cellsThe cortical microtubules were arranged into a fine structure with a predominant orientation parallel to the short axis of the cells in the control cells. The pattern of microtubules changed noticeably when cells were subjected to hyperosmotic stress. Denser cortical microtubule arrays were observed in osmotic-stressed cells than in control cells (Fig. 1). The ABA content also increased gradually with extension of the treatment time when maize roots were subjected to osmotic stress caused by 15% PEG-6000 (Fig. 2).
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Effects of PEG-6000, oryzalin, and taxol treatments on
-tubulin arrangement of maize root cellsNormally cell microtubules in the epidermal cells of maize roots are organized in parallel, straight, and dense parallel bundles (Fig. 3). After dehydration (PEG-6000 treatment), the microtubule organization was markedly altered and the fine microtubule structure disappeared whereas some thicker structures formed (Fig. 3C). The effects of microtubule-destabilizing and -stabilizing drugs on microtubule arrangement in maize radicular cells were also analysed. As expected, no microtubules were visible after oryzalin treatment (Fig. 3E). When the cells were treated with taxol, the microtubule arrays were thicker than those of the control (Fig. 3G).
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The roles of microtubule arrangement in ABA accumulation induced by osmotic stress in maize roots
The above results provided evidence that microtubules displayed cellular responses to the osmotic stress in maize roots, and osmotic stress induced a significant accumulation of ABA. However, it is still unclear whether there is a relationship between the dynamic changes to microtubules and ABA accumulation under osmotic stress conditions. Therefore, the effects of the microtubule-disrupting agent oryzalin on the microtubule arrangement in the radicular cells during dehydration were studied. The results from immunofluorescent staining showed that microtubule arrays were broken down into smaller fragments in all oryzalin treatments (Fig. 4). However, compared with the cells treated with oryzalin alone (Fig. 4C), the
-tubulin arrays could be partially recovered when oryzalin-treated cells were then subjected to osmotic stress treatment, i.e. some new -tubulin arrays were re-formed (Fig. 4E). No such recovery of -tubulin arrays was observed when osmotic stress-pretreated cells were treated with oryzalin (Fig. 4G). These results support the notion that the dynamic changes of microtubules were involved in the desiccation response in Z. mays.
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Figure 5A shows the effects of different treatments on ABA accumulation in maize root cells. PEG treatment significantly stimulated biosynthesis and accumulation of ABA, as compared with the control. Both oryzalin treatment and PEG treatment after pretreatment with oryzalin doubled the ABA content, which was much less than that of root cells treated with PEG alone. However, the ABA content of cells treated with oryzalin after being pretreated with PEG was significantly higher than those of other treatments.
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In order to explore the possible mechanism(s) of the effects of different treatments on ABA biosynthesis in maize roots, the effects of these treatments on the expression of the Vp14 gene that encodes a key regulatory enzyme catalysing ABA biosynthesis, 9-cis-epoxycarotenoid dioxygenase (Chernys and Zeevaart, 2000), were investigated using RT-PCR (Fig. 5B). The results indicated that all treatments had various degrees of stimulating effects on the expression of the Vp14 gene, as compared with the control treatment, especially for treatments with PEG and PEG/oryzalin.
The effects of the microtubule-stabilizing agent taxol on -tubulin dynamics of maize root cells during the dehydration process were also investigated. Compared with control (no taxol or PEG treatment), the -tubulin became more stable and microtubules became thicker after taxol treatment alone (Fig. 6C). However, when the taxol-treated cells were further exposed to osmotic stress, the direction of the microtubule arrays became disordered (Fig. 6E). When root cells were first treated with PEG then with taxol, the microtubules became thicker, but the direction of the microtubule arrays was altered (Fig. 6G)
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Similarly, the biosynthesis and accumulation of ABA in root cells were also significantly affected by the treatments with either taxol, PEG, or both (Fig. 7A). When root cells were exposed to either taxol or PEG alone, or combined treatment with taxol and PEG, the ABA content of these cells increased in comparison with that of control (Fig. 7). However, significant differences in the extent of this stimulation of ABA biosynthesis existed among different treatments. The ABA contents of the cells treated with either PEG or taxol increased around 3-fold and 2-fold, respectively. The effects of PEG and taxol combined treatment on the ABA content of cells varied greatly depending on the order of treatments. When cells were treated with taxol after PEG, their ABA content was 6–7 times higher than in the control cells, whereas no obvious difference in ABA content of the cells was observed when treated with taxol prior to PEG in comparison with those treated with either PEG or taxol alone (Fig. 7A). RT-PCR results showed that the expression of the Vp14 gene was obviously stimulated by either PEG treatment or taxol treatment, or both, as compared with the control (Fig. 7B).
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Discussion |
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The plant cytoskeleton readily remodels in response to various intracellular and external stimuli. It is not surprising that the dynamic behaviour of microtubules, as one of the most important components of the cytoskeleton, is considered to be responsible for these stimuli-induced responses and thus regulates cell growth and morphogenesis (Whittington et al., 2001; Komis et al., 2002; Dixit and Cyr, 2004).
In the last two decades, our understanding of the processes underlying plant responses to drought, at both the molecular and whole-plant level, has rapidly progressed. When exposed to drought stress, plants exhibit a wide range of responses, from physiological to molecular events (Zhang et al., 2004, 2006). At the level of physiological processes, the rapid biosynthesis and significant accumulation of ABA in plant cells is the earliest and most obvious physiological event upon drought stress (Liang and Zhang, 1996, 1997a, b), perhaps largely due to the increased expression of genes related to ABA biosynthesis (Chernys and Zeevaart, 2000; Seo and Koshiba, 2002). ABA is considered to be a major phytohormone that can enhance plant tolerance to drought, either through controlling cell water status or through stimulating gene expression and cell membrane stability (Bray, 2004). Therefore, it is vitally important to explore the mechanisms of drought-induced ABA biosynthesis (Zhang et al., 2006). Unfortunately, little is known about them so far, especially the upstream components of drought signalling that induces ABA biosynthesis, for example, how plant cells perceive the drought signal.
Several pieces of evidence showed that the relaxation of the plasma membrane or shrinkage of cell volume may be related to the osmotic stress-induced ABA biosynthesis (Creelman and Mullet, 1991; Jia et al., 2001). However, it is still unclear how drought stress affects the cell wall–plasma membrane interaction and how this interaction triggers the biosynthesis of ABA. Previous studies have provided biochemical and immunological evidence that integrin-like proteins mediate the interaction between cell wall and plasma membrane and are involved in the drought stress-induced ABA biosynthesis (Lü et al., 2007b). Further results showed that a strong interaction exists between integrin-like proteins and -tubulin protein (Lü et al., 2007a). It is reasonable to assume that there is a cell wall–plasma membrane–cytoskeleton continuum in plant cells, which may be responsible for the osmotic stress-induced ABA biosynthesis.
To test this hypothesis, the possible roles of microtubule dynamic changes (i.e. polymerization and depolymerization) in osmotic stress-induced ABA biosynthesis were investigated. A lot of evidence has shown that, under normal conditions, microtubules constantly remodel their arrangement in order to adjust cell growth. Therefore, any factor causing damage to the microtubule dynamic balance may affect physiological processes, and thus cell growth. The results presented here showed that microtubule organization in radicular cells of maize roots is responsive to osmotic stress, and osmotic stress altered the dynamic balance of -tubulin proteins from depolymerization to polymerization, and, as a result, denser and brighter cortical microtubule arrays were visualized in root cells when subjected to osmotic stress (Fig. 1). Significant accumulation of ABA was also observed in maize roots when exposed to the same osmotic stress conditions (Fig. 2). In order to explore the possible roles of microtubule dynamics in osmotic stress-induced ABA accumulation, the effects of two microtubule-specific drugs, oryzalin and taxol, on microtubular behaviour and ABA content in maize root cells under either control (no PEG treatment) or osmotic stress (PEG treatment) conditions were examined. The results showed that approximately all microtubules disappeared from root cells when treated with oryzalin, a drug which disrupts the microtubular cytoskeleton (Fig. 3). The effect of oryzalin on microtubule organization would depend on the treatment time when this was combined with osmotic stress, and partial recovery of microtubule arrays was visualized when root cells were exposed to osmotic stress after oryzalin treatment. However, more serious depolymerization of microtubule arrays was detected when oryzalin was applied after osmotic stress treatment (Fig. 4). Unfortunately, little is known about the mechanism of interaction between oryzalin and osmotic stress in microtubule organization. The effects of osmotic stress on taxol-stabilized microtubule arrays also varied with the treatment sequence of taxol, but only different extents of orientation of microtubule arrays were observed (Fig. 6). Similarly, it is unclear why osmotic stress altered the orientation of microtubule arrays in these cases.
The results in this experiment and previous results have shown that microtubule dynamics are responsive to osmotic stress, and, moreover, microtubules interact with integrin-like proteins that have been shown to be involved in cell wall–plasma membrane interaction and osmotic stress-induced ABA accumulation (Lü et al., 2007a). In the present study, further evidence has been provided that microtubule dynamics may be involved in osmotic stress-induced ABA accumulation. Pharmacological approaches that altered microtubule stability were used to analyse ABA biosynthesis under osmotic stress conditions. Both disruption and stabilization of microtubules significantly stimulated ABA accumulation in maize root cells, although these stimulations were markedly lower than that produced by osmotic stress treatment (Figs 5, 7). Furthermore, cells with changed microtubular stability did not respond further to osmotic stress in terms of ABA biosynthesis. However, treatments with both a microtubule destabilizer and a stabilizer enhanced the sensitivity of cells to osmotic stress in terms of ABA accumulation. It is suggested that both osmotic stress and changes in microtubule dynamics would trigger maize root cells to biosynthesize ABA, and interactions between osmotic stress and microtubule dynamics would have an effect on ABA accumulation in root cells, although the exact mechanism is not clear at present. However, it may involve the regulation of gene expression at the transcriptional level (Figs 5B, 7B).
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Acknowledgements |
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The authors thank Professor Ming Yuan (College of Biological Sciences, China Agricultural University, Beijing, China) for his help in
-tubulin immunofluorescence staining. This work was supported by the National Natural Science Foundation of China (grant no. 30471046), State Key Basic Research and Development Program of China (grant no. 2003CB114303), and Natural Science Foundation of Jiangsu Province (grant no. BK2002048). |
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