| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
|
First published online June 1, 2004; 10.1104/pp.103.032631 Plant Physiology 135:849-858 (2004) © 2004 American Society of Plant Biologists
Plastidial
|
|||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
 |
ABSTRACT |
|---|
 TOP ABSTRACT RESULTSDISCUSSIONMATERIALS AND METHODSLITERATURE CITED |
|---|
-glucan phosphorylase in starch metabolism in the leaves of Arabidopsis, two independent mutant lines containing T-DNA insertions within the phosphorylase gene were identified. Both insertions eliminate the activity of the plastidial -glucan phosphorylase. Measurement of other enzymes of starch metabolism reveals only minor changes compared with the wild type. The loss of plastidial -glucan phosphorylase does not cause a significant change in the total accumulation of starch during the day or its remobilization at night. Starch structure and composition are unaltered. However, mutant plants display lesions on their leaves that are not seen on wild-type plants, and mesophyll cells bordering the lesions accumulate high levels of starch. Lesion formation is abolished by growing plants under 100% humidity in still air, but subsequent transfer to circulating air with lower humidity causes extensive wilting in the mutant leaves. Wilted sectors die, causing large lesions that are bordered by starch-accumulating cells. Similar lesions are caused by the application of acute salt stress to mature plants. We conclude that plastidial phosphorylase is not required for the degradation of starch, but that it plays a role in the capacity of the leaf lamina to endure a transient water deficit.
-Glucan phosphorylase (EC 2.4.1.1) is a key enzyme in glucan catabolism in animals, fungi, and prokaryotes (Newgard et al., 1989
). It is also considered to be an important enzyme in the degradation of starch in plants, but definitive evidence for this role is lacking (Preiss, 1982; Beck and Ziegler, 1989; Sonnewald et al., 1995; Trethewey and Smith, 2000). The enzyme catalyzes the phosphorolysis of the terminal residue from the nonreducing ends of -1,4-linked glucan chains, liberating Glc-1-phosphate (Glc-1-P). Although this reaction is reversible, it has been argued that the relatively low levels of Glc-1-P and high levels of inorganic phosphate in plant cells favor the glucan-degrading reaction (Kruger and ap Rees, 1983a).
All plants studied so far have plastidial and cytosolic isoforms of phosphorylase, which are encoded by separate genes (Nakano et al., 1989; Mori et al., 1991; Van Berkel et al., 1991; Buchner et al., 1996). Starch is synthesized exclusively in plastids, so only the plastidial isoform is implicated in its metabolism. The amino acid sequence of -glucan phosphorylase is conserved between prokaryotes and eukaryotes (Newgard et al., 1989). However, plastidial isoforms of starch phosphorylase differ significantly from all other forms due to the presence of an insert of 80 amino acids in length. This domain contributes toward the marked difference in substrate preference between the plastidial and cytosolic isoforms (Mori et al., 1993). The plastidial form (L-form) has a low affinity for large branched glucans and a high affinity for small linear maltodextrins while the cytosolic isoform (H-form) has the opposite relative affinities.
Chloroplasts contain amylases in addition to phosphorylase (Stitt et al., 1978; Okita et al., 1979; Lin et al., 1988; Li et al., 1992). Starch degradation in isolated chloroplasts results in either phosphorylated or nonphosphorylated products or both depending upon incubation conditions (Stitt and Heldt, 1981; Kruger and ap Rees, 1983b). However, the relative contributions of the phosphorolytic and hydrolytic pathways in vivo are not known. Stitt et al. (1985) suggested that the products of phosphorolysis and hydrolysis could have different metabolic fates, supporting respiration and Suc synthesis, respectively. Antisense repression of a gene encoding a plastidial isoform of phosphorylase in potato (Solanum tuberosum) decreased the detectable phosphorylase activity in leaves but had no major impact on the accumulation of starch (Sonnewald et al., 1995). However, the nature of the antisense technique and the fact that potato contains a second plastidial isoform, which is expressed in leaves (Albrecht et al., 2001), mean that an important function for phosphorylase in starch metabolism could not be ruled out.
The availability of knockout mutants in Arabidopsis now allows us to explore the precise role of plastidial -glucan phosphorylase in starch degradation and synthesis. Here we report that complete loss of the enzyme does not cause a significant change in the overall accumulation of starch during the day, or its remobilization at night in healthy plants. However, we present evidence that phosphorylase-deficient plants are more sensitive to transient water and salt stress and speculate that, by providing substrates for chloroplast respiratory metabolism, the phosphorolysis of starch may play an important role in stress tolerance.
|
RESULTS |
|---|
|
TOPABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSLITERATURE CITED |
|---|
-Glucan Phosphorylase Mutants
The Arabidopsis genome contains two genes on chromosome 3 encoding isoforms of -glucan phosphorylase (At3g29320 and At3g46970). Full-length cDNAs (GenBank accession nos. AY049235 and BT003012, respectively) confirm the gene predictions. The protein encoded by At3g46970 is 841 amino acids long and does not have a predicted chloroplast transit peptide. This most likely encodes a cytosolic isoform (AtPHS2). The protein encoded by At3g29320 is 962 amino acids long including a domain of approximately 80 amino acids characteristic of plastidial isoforms of phosphorylase, and a putative amino-terminal chloroplast transit peptide of 62 amino acids (ChloroP; Emanuelsson et al., 1999). This indicates that At3g29320 encodes the plastidial isoform of -glucan phosphorylase (AtPHS1). Two Arabidopsis lines containing T-DNA insertions in the AtPHS1 gene were identified from the collection produced at INRA Versailles (Bechtold et al., 1993; Bouchez et al., 1993). The first (Atphs1-1) was identified by screening DNA pools by PCR using multiple primer combinations (Krysan et al., 1996). The second (Atphs1-2) was identified by the Genoplante FLAGdb/FST initiative.
T-DNA-specific PCR primers close to the left border gave products with AtPHS1-specific primers for both Atphs1-1 and Atphs1-2. Sequence analysis revealed the precise location of each T-DNA insertion within the AtPHS1 gene (Fig. 1). We used PCR to identify plants that were homozygous for each T-DNA insertion. The disruption of the AtPHS1 gene in Atphs1-1 is expected to result in a null mutation. Reverse transcription (RT)-PCR analysis confirmed that no functional AtPHS1 mRNA is produced, although hybrid transcripts containing both AtPHS1 and T-DNA sequences are produced (not shown). In Atphs1-2, the insertion site is upstream of the ATG translational start codon.RT-PCR and RNA gel-blot analyses established that this allele is transcribed from a promoter within theT-DNA, producing a transcript larger than AtPHS1 mRNA (not shown).
|
We used biochemical assays and native PAGE techniques to analyze the activity of phosphorylase and other starch metabolizing enzymes in wild-type and homozygous Atphs1 plants. Enzymes were assayed under conditions previously determined to be optimal, giving activity that was linear with respect to time and volume of extract added (Zeeman et al., 1998a; Critchley et al., 2001). Phosphorylase was measured by the generation of Glc-1-P from a glucan substrate and Pi. Total activity was significantly lower in Atphs1-1 than in the wild type, using three different glucan substrates (amylopectin, glycogen, and maltoheptaose). The difference was most pronounced using maltoheptaose and least pronounced using glycogen (Table I). This is consistent with a reduction in plastidial phosphorylase. The plastidial isoform from other species has a marked preference for short linear malto-oligosaccharides over large branched substrates, whereas the converse is true for the cytosolic isoform (Shimomura et al., 1982; Steup, 1988).
|
). Wild-type plants contained both forms of phosphorylase, but Atphs1-1 and Atphs1-2 contained no detectable plastidial activity (Fig. 2). We conclude that although the coding region is not disrupted in Atphs1-2, the aberrant transcript produced from this gene is either not translated, or results in greatly reduced amounts of AtPHS1 protein, or produces an aberrant protein.
|
-Amylase, pullulanase, maltase, starch synthase, and starch branching enzyme were unchanged in Atphs1-1 relative to the wild type. -Amylase activity was 60% higher and disproportionating enzyme 30% lower in the mutant relative to the wild type (Table I). Activity gels were also used to analyze the isoforms of starch hydrolyzing enzymes, starch synthases, and starch branching enzymes. No differences between wild type and mutants were observed (not shown).
Loss of Plastidial Phosphorylase Does Not Alter Total Starch Content or Starch Structure
We measured the starch content in wild type and Atphs1-1 leaves over the diurnal cycle. No difference between wild-type and mutant plants was observed (Fig. 3A). There was also no difference in the distribution of chain lengths of the amylopectin fraction (Fig. 3B), or in the amylose to amylopectin ratio of starch extracted from leaves at the end of the day (Fig. 3C). Comparison of the Suc and free hexose contents of the leaves over the diurnal cycle revealed only minor differences between the wild type and mutant (Fig. 3A). The results indicate that under the growth conditions used, the loss of phosphorylase has little or no impact on the overall starch and sugar metabolism in the leaves.
|
We observed a consistent phenotypic difference between wild type and both Atphs1 lines. The leaves of mature mutant plants consistently had small, white lesions on the tips or margins of fully expanded leaves. Both independent mutant lines displayed the lesion-forming phenotype and in each line, lesion formation cosegregated with the loss of plastidial phosphorylase. The lesions did not increase in size after appearance and were not bordered by chlorotic tissue (Fig. 4A). The frequency and severity of lesions varied between batches of plants, but lesion-free mutant plants were rare. Further examination of the lesions on leaves of the mutant plants revealed that high levels of starch were present in the living cells bordering each lesion at the end of the night, when the rest of the leaf had metabolized its starch (Fig. 4B). This starch accumulation occurred in a highly cell-specific manner (Fig. 4C) with adjacent starch-rich and starch-free cells. At the end of the day, all cells contained starch and could not be distinguished by qualitative iodine staining (not shown). Because the regions of starch accumulation were so small compared with the total leaf area, the measurements of total leaf starch (Fig. 3A) did not detect this difference between the wild type and mutant.
|
-amylase and D-enzyme (Table I) were altered specifically in lesion-bearing leaves. A separate batch of plants was grown and -amylase andD-enzyme were measured in wild-type leaves and in Atphs1-1 leaves that were either healthy or that contained visible lesions. The results (Table II) indicate that the changes in these enzymes are associated with lesion formation. The activities were the same in wild-type leaves and mutant leaves without lesions, but in mutant leaves with lesions, the activity of -amylase was increased 80% and that of D-enzyme was decreased 50%, relative to the wild type.
|
We observed that the growth conditions influenced the occurrence of lesions on the mutant plants. A particularly high frequency was observed after seedlings had been transplanted from germination medium to individual pots. After transplantation, trays of individual pots were well watered and covered for 5 d with a clear plastic lid to increase the humidity near to 100%. Lesions developed on the mutant leaves over a period of 3 to 5 d after the lid was removed.
We reasoned that the sudden reduction in humidity might be promoting lesion formation. Accordingly, plants were grown from seed at close to 100% humidity in still air for 5 weeks. Transplantation to potting compost was carried out after 3 weeks. No lesions were visible at 3 weeks or at 5 weeks on either wild-type or mutant plants (Fig. 5A). After 5 weeks, the plants were shifted abruptly to circulating air with 60% ± 5% relative humidity at the end of their normal photoperiod, then photographed and scored for lesions on successive days. After 1 d the fully expanded leaves of the mutant plants were severely wilted. Wilted sectors died, leading to lesions similar to those seen under normal growth conditions but more severe (Fig. 5, B and C). Minor symptoms were visible on some fully expanded wild-type leaves. Leaves present at the time of the shift to lower humidity were scored as containing no lesion, minor lesion (
30% leaf area), major lesion (
30% leaf area), and dead. In the mutant, 75% of the leaves developed lesions, the majority of which were severe. In the wild type only 25% of the leaves developed lesions, very few of which were severe (Fig. 6). Lesion-bearing leaves were stained with iodine 6 d after the shift to lower humidity. Starch accumulating cells were visible bordering the lesions in the mutant plants, but not in the wild type (not shown). Leaves of Atphs1-1 that emerged after the shift to lower humidity exhibited far fewer lesions than the mature leaves. No lesions were visible on new wild-type leaves (Fig. 5B). A second batch of plants treated identically gave very similar results.
|
|
We investigated the stomatal response to an abrupt exposure to water deficit. First, we detached mutant and wild-type rosettes from their root systems and weighed them at intervals over 60 min. There was appreciable water loss at first, but the rate rapidly decreased and was 20% to 25% of the initial value after 60 min (Fig. 7A). The rate of water loss was the same from both mutants and wild-type plants. Second, we investigated gas exchange of individual mutant and wild-type leaves using infrared gas analysis. Leaves that had attained a steady state of gas exchange in the cuvette were detached to stop their water supply. This caused an appreciable and essentially identical decrease in stomatal conductance in mutant and wild type plants (Fig. 7B). Abruptly decreasing the relative humidity of the air around attached leaves by 75% resulted in increased transpiration, but not stomatal closure, in both the mutants and in the wild type (not shown). Subsequent detachment of the leaf did cause stomatal closure. We used light microscopy to examine the starch content of guard cells in iodine-stained epidermal peels. There was no visible difference between the wild type and Atphs1-1 in the starch content of guard cell chloroplasts 1 h before the start and 1 h before the end of the photoperiod. Both lines contained starch at both time points (not shown). Together, these results suggest that stomatal function in the mutants does not differ from that of the wild type.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSLITERATURE CITED |
|---|
Our results show that the loss of plastidial -glucan phosphorylase does not impact significantly on the metabolism of starch when viewed on a whole plant basis. Leaves of two independently isolated knockout mutants accumulate and degrade the same amount of starch as do wild-type plants. There are no detectable alterations in the amylopectin structure or the composition of the starch, no major changes in the activities of other starch-metabolizing enzymes, and the contents of Suc and free hexoses do not differ appreciably from the wild type. If starch phosphorylase catalyzes a major degradative (or synthetic) flux in vivo, our results indicate that it can be replaced in this role, and is thus redundant. Although this result is surprising given the frequent suggestions that phosphorylase is important in starch degradation in leaves (Preiss, 1982; Beck and Ziegler, 1989; Trethewey and Smith, 2000), it is consistent with a growing body of evidence to the contrary. The activity of plastidial phosphorylase in Arabidopsis is low when compared with the amylolytic activities and alone is insufficient to catalyze the observed rate of starch degradation (Lin et al., 1988; Zeeman et al., 1998a). Furthermore, reduction of the plastidial isoforms of phosphorylase in potato using antisense techniques had no reported effects on leaf starch metabolism (Sonnewald et al., 1995; also see introduction). Thus, we conclude that in Arabidopsis, as may also be the case in potato, starch degradation in mesophyll chloroplasts is primarily hydrolytic.
The Role of Plastidial Phosphorylase
Although analysis of total leaf starch and sugar contents revealed no differences between mutant and wild-type plants, the mutants did display a consistent lesion-forming phenotype, with a marked accumulation of starch around the lesion sites. This starch accumulation was very localized and would not contribute significantly to the whole plant measurements. Lesions were observed in both of the independently isolated Atphs1 mutants and cosegregated with the loss of phosphorylase. The localized starch accumulation is consistent with the mutant phenotypes resulting directly from disruption of the Atphs1 gene, rather than from a secondary effect of T-DNA insertion in the AtPHS1 gene affecting neighboring genes. The failure to mobilize starch via phosphorylase in certain cells under specific environmental conditions may result in cell death and be the basis of the lesion formation.
By altering the growth conditions and imposing salt stress we have provided evidence that the resilience of Arabidopsis plants to acute water stress conditions is compromised in Atphs1 mutants. Lesion formation was not observed when plants were grown in still air with high humidity, but was strongly promoted by transfer to low humidity with circulating air and by treatment of the roots with a salt solution. Both changes in conditions are likely to cause a rapid transient water stress in leaves. The combination of still air and high humidity would promote a high stomatal conductance. A sudden increase in transpiration rate due to air circulation and a drop in humidity could result in the loss of excessive amounts of water, particularly from the tips and margins of the leaves, which represent the extremities of the transpirational path. Indeed, the consequence of this stress was visible on the wild-type plants as the margins of some of the largest, mature leaves wilted and died after the change in conditions. The acute salt treatment would also result in the loss of excessive water from the leaf lamina, but in this case through the reduced uptake into the roots and transpirational flow. In addition this treatment would cause a long-term stress as the plant takes up salt.
We suggest two possible reasons for the increased sensitivity of Atphs1 plants to these conditions. First, the loss of plastidial phosphorylase could affect stomatal function such that transpirational water loss from the mutant leaves is greater than from the wild type, causing indirect symptoms in the leaf lamina. The metabolism of starch in guard cells has long been viewed as an important source of carbohydrate for potassium counter-ion synthesis or sugar accumulation during stomatal opening (Lloyd, 1908; Outlaw and Manchester, 1979). However, we do not favor this hypothesis because a block in starch degradation would result in a failure of stomatal opening rather than a failure of closure. Furthermore, our results indicate that stomatal closure occurs at the same rate in the mutant as in the wild type (Fig. 7). Second, the loss of plastidial phosphorylase could compromise the ability of the mutant palisade and mesophyll cells to tolerate the transient water stress, leading to cell death and wilting. Such a role is consistent with the local accumulation of starch around the lesion sites. We suggest that in these cells, a shift away from starch hydrolysis toward phosphorolysis has been triggered. The failure of the phosphorolytic pathway in these conditions could thus cause both lesion formation and local starch accumulation. Although we favor this second hypothesis, at this stage we cannot exclude the possibility that lesion phenotype is pleiotropic and only indirectly linked to the loss of plastidial phosphorylase.
The Role of Hexose Phosphates Supplied by Phosphorylase
The role of phosphorylase may be to provide hexose phosphates as substrates for the oxidative pentose phosphate pathway (OPPP) inside the chloroplast at night. This pathway utilizes Glc-6-P to provide reducing power (NADPH) for many biosynthetic reactions but is also crucial for controlling the levels of reactive oxygen intermediates via the ascorbate-glutathione cycle, thereby avoiding oxidative stress. Reactive oxygen intermediates are produced from multiple sources in the cell during abiotic stress conditions (including water deficit) and can diffuse between cellular compartments (Smirnoff, 1993; Mittler, 2002). By limiting the operation of the chloroplast OPPP, the loss of phosphorylase could also limit the scavenging of reactive oxygen species. Mesophyll chloroplasts are not able to obtain hexose-phosphate from the cytosol, as they lack the ability to translocate hexose-phosphates across the inner envelope (Fliege et al., 1978). Furthermore, chloroplasts are reported to lack hexokinase preventing the conversion of Glc to Glc-6-P (Stitt et al., 1978; Wiese et al., 1999), although this point has recently been questioned (Olsson et al., 2003). The import of five-carbon sugars via the recently discovered xylulose5-P/Pi exchange transporter (Eicks et al., 2002) could potentially provide substrates for the OPPP. We speculate that when the demand for reducing power in the chloroplast at night is high or is suddenly increased by imposition of conditions that generate an oxidative stress, direct provision of hexose-phosphate within the chloroplast via phosphorolytic starch degradation may be essential. Other pathways that could supply substrates for the OPPP may be inadequate. The hypothesis that the lesions on leaves of plants lacking plastidial phosphorylase are a consequence of impaired functioning of chloroplastic OPPP in the dark requires further investigation.
We observed that the sensitivity of the mutants to water stress was restricted to existing mature leaves. New leaves that developed after the application of stress conditions did not develop lesions to the same extent, indicating that adaptive changes occurred to enhance tolerance to the new conditions. Thus, phosphorylase probably acts to provide a very rapid response to a sudden environmental change, and is followed by altered gene expression for complete acclimation. Interestingly, a hexose-phosphate transporter that could provide substrates for the oxidative pentose phosphate pathway in the dark is induced by water stress in spinach leaves (Quick et al., 1995). The induction of additional tolerance mechanisms could explain why the penetrance of the lesion-forming phenotype was dependent on the precise growth conditions and why lesion-bearing phenotypes have not been reported for starchless mutants. Producing double mutants lacking both starch and phosphorylase will provide a means to test this theory.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSLITERATURE CITED |
|---|
A population of T-DNA-transformed Arabidopsis (Wassilewskija ecotype) was obtained from Institut National de la Recherche Agronomique-Versailles. DNA was prepared from pools of 100 lines in collaboration with P. Benoist and M. Thomas, University of Paris-Sud, Orsay, France. PCR screening was carried out as described by Germain et al. (2001) and allele Atphs1-1 found in pool15/5. The mutant line 125B01 was obtained from the Flagdb/FST initiative (http://flagdb-genoplante-info.infobiogen.fr/projects/fst/) and designated Atphs1-2.
Growth Conditions and Gas Exchange Measurements
Plants were grown in a greenhouse or a controlled environment chamber. Unless otherwise stated the conditions were as follows. Greenhouse temperature was maintained above 10°C and natural illumination supplemented to provide a minimum photoperiod of 16 h. The controlled environment chamber provided a constant 20°C, 75% relative humidity, and a 12-h/12-h light/dark cycle, with uniform illumination of 175 µmol photons m–2 s–1. Sown seeds were covered with a clear plastic lid and stratified (4°C, 2 d). Lids were removed 10 d after sowing when the cotyledons had emerged. Seeds were sown either directly onto a peat-based potting compost or germinated first on fine grade seed compost, then transplanted after 2 to 3 weeks into individual pots (5 x 5 x 5 cm) containing potting compost. Transplanted seedlings were again covered with a clear plastic lid for 5 d. To administer acute salt stress, pots containing mature rosettes were immersed in a solution of 500 mM NaCl for 15 min. The rosettes were kept dry. Excess solution was drained and the application repeated twice, each time with fresh salt solution. Plants were further watered on subsequent days with salt solution. To determine kanamycin resistance, seeds were germinated on 0.7% (w/v) agar plates containing 50 mg/L kanamycin and the seedlings scored for resistance after 2 weeks.
Gas exchange measurements were made using a CIRAS 1 infrared gas analyser (PP Systems, Hitchin, UK). Mature leaves were placed in the cuvette and illuminated using lights of the growth cabinet. When the leaves had equilibrated, achieving a steady state of gas exchange, the petioles were cut and changes in gas exchange measured over the course of 1 h.
Materials
All enzymes and biochemicals were from Roche (Poole, Dorset, UK) except Taq polymerase, isoamylase, and starch azure, which were from Sigma-Aldrich Chemical (Poole, Dorset, UK).
Native PAGE
Native PAGE for isoforms of -glucan phosphorylase was adapted from Steup (1990). Leaves harvested midway through their photoperiod were extracted in 100 mM 3-[N-morpholino] propanesulfonic acid), pH 7.2, 1 mM dithiothreitol, 1 mM EDTA, and 10% (v/v) ethanediol (extraction buffer). Gels were run exactly as described previously (Zeeman et al., 1998a) except that the incubation medium contained 100 mM Tris-HCl, pH 7.0, 1 mM dithiothreitol, and 20 mM Glc-1-P. Native PAGE for the detection of starch hydrolyzing activities and branching enzymes isoforms were performed exactly as described previously (Zeeman et al., 1998b). Native PAGE for the detection of starch synthases was performed as described by Edwards et al. (1995).
Enzyme Measurements
Enzyme assays were previously optimized with respect to pH and the concentrations of all components of the assays using extracts of leaves of the Wassilewskija wild-type ecotype (Zeeman et al., 1998a; Critchley et al., 2001). -Amylase, -amylase, maltase, disproportionating enzymes, starch synthase, and starch branching enzyme were measured as described in Zeeman et al. (1998a). -Glucan phosphorylase was assayed using a continuous assay in the direction of Glc-1-P formation, coupled to the production of NADH. Leaves were extracted in extraction buffer (see above). The 1-mL reaction mixture contained 20 mM 3-[N-morpholino] propanesulfonic acid), pH 7.0, 20 mM Na2HPO4/KH2PO4, 10 mM MgCl2, 3.4 mM NAD, 1 unit phosphoglucomutase (from rabbit muscle), 1 unit Glc-6-phosphate dehydrogenase (from Leuconostoc mesenteroides), 2.5 µM Glc-1,6-bisphosphate, and the glucan substrate (amylopectin, glycogen, or maltoheptaose, with final concentrations of 2.5 mg mL–1, 1.0 mg mL–1, and 1 mM, respectively).
Metabolite Measurements
For the extraction and measurements of starch and sugars, samples comprising all the leaves of individual plants were harvested and immediately frozen in liquid N2. Extracts were made using 0.7 M perchloric acid as described in Critchley et al. (2001). Starch was measured in the insoluble pellet (Critchley et al., 2001). Suc and hexoses were measured as described in Zeeman and ap Rees (1999). Leaves were stained for starch by killing and decolorizing the tissue in hot 80% ethanol, then staining with Lugols solution (Sigma).
Starch Composition and Structure
Separation of amylose and amylopectin using Sepharose CL2B (Sigma) in a 9-mL column was performed as described by Denyer et al. (1995). The distribution of chain lengths of amylopectin was analyzed using the fluorophore assisted PAGE system described by Edwards et al. (1999).
Sequence data from this article have been deposited with the EMBL/GenBank data libraries under accession numbers AP001309 and AL133292.
|
ACKNOWLEDGMENTS |
|---|
Received August 31, 2003; returned for revision March 10, 2004; accepted March 10, 2004.
|
FOOTNOTES |
|---|
Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.103.032631.
* Corresponding author; e-mail sam.zeeman{at}ips.unibe.ch; fax 41–31–631–5222.
|
LITERATURE CITED |
|---|
|
TOPABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSLITERATURE CITED |
|---|
Bechtold N, Ellis J, Pelletier G (1993) In planta Agrobacterium-mediated gene transfer by infiltration of adult Arabidopsis thaliana plants. C R Acad Sci Paris Life Sci 316: 1194–1199
Beck E, Ziegler P (1989) Biosynthesis and degradation of starch in higher plants. Annu Rev Plant Physiol Plant Mol Biol 40: 95–117[CrossRef][Web of Science]
Bouchez D, Camilleri C, Caboche M (1993) A binary vector based on Basta resistance for in planta transformation of Arabidopsis thaliana. C R Acad Sci Paris Life Sci 316: 1188–1193
Buchner P, Borisjuk L, Wobus U (1996) Glucan phosphorylases in Vicia faba L.: cloning, structural analysis and expression patterns of cytosolic and plastidic forms in relation to starch. Planta 199: 64–73[Web of Science][Medline]
Critchley JH, Zeeman SC, Takaha T, Smith AM, Smith SM (2001) A critical role for disproportionating enzyme in starch breakdown is revealed by a knock-out mutation in Arabidopsis. Plant J 26: 89–100[CrossRef][Web of Science][Medline]
Denyer K, Barber LM, Burton R, Hedley CL, Hylton CM, Johnson S, Jones DA, Marshall J, Smith AM, Tatge H, et al (1995) The isolation and characterisation of novel low amylose mutants of Pisum sativum L. Plant Cell Environ 18: 1019–1026[CrossRef]
Edwards A, Fulton DC, Hylton CM, Jobling SA, Gidley M, Rossner U, Martin C, Smith AM (1999) A combined reduction in activity of starch synthases II and III of potato has novel effects on the starch of tubers. Plant J 17: 251–261[CrossRef][Web of Science]
Edwards A, Marshall J, Sidebottom C, Visser RGF, Smith AM, Martin C (1995) Biochemical and molecular characterisation of a novel starch synthase from potato tubers. Plant J 8: 283–294[CrossRef][Web of Science][Medline]
Eicks M, MaurinoV, Knappe S, Flügge U-I, Fischer K (2002) The plastidic pentose phopsphate translocator represents a link between the cytosolic and the plastidic pentose phosphate pathways in plants. Plant Physiol 128: 512–522
Emanuelsson O, Nielsen H, von Heijne G (1999) ChloroP, a neural network-based method for predicting chloroplast transit peptides and their cleavage sites. Protein Sci 8: 978–984[Web of Science][Medline]
Fliege R, Flügge UI, Werdan K, Heldt HW (1978) Specific transport of inorganic-phosphate, 3-phosphoglycerate and triosephosphates across inner membrane of envelope in spinach-chloroplasts. Biochim Biophys Acta 502: 232–247[Medline]
Germain V, Rylott E, Larson TR, Sherson SM, Bechtold N, Carde J-P, Bryce JH, Graham IA, Smith SM (2001) Requirement for 3-ketoacyl-CoA thiolase-2 in peroxisome development, fatty acid -oxidation and breakdown of triacylglycerol in lipid bodies of Arabidopsis seedlings. Plant J 28: 1–12[CrossRef][Web of Science][Medline]
Kruger NJ, ap Rees T (1983a) Properties of -glucan phosphorylase from pea. Phytochem 22: 1891–1898[CrossRef]
Kruger NJ, ap Rees T (1983b) Maltose metabolism by pea chloroplasts. Planta 158: 179–184[CrossRef]
Krysan PJ, Young JC, Tax F, Sussman MR (1996) Identification of transferred DNA insertions within Arabidopsis genes involved in signal transduction and ion transport. Proc Natl Acad Sci USA 93: 8145–8150
Li B, Servaites JC, Geiger DR (1992) Characterization and subcellular localization of debranching enzyme and endoamylase from leaves of sugar-beet. Plant Physiol 98: 1277–1284
Lin T-P, Spilatro SR, Preiss J (1988) Subcellular localization and characterization of amylases in Arabidopsis leaf. Plant Physiol 86: 251–259
Lloyd FE (1908) The behaviour of stomata. Carnegie Inst Washington Publ 82: 1–142
Mittler R (2002) Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci 7: 405–410[CrossRef][Web of Science][Medline]
Mori H, Tanizawa K, Fukui T (1991) Potato tuber type H phosphorylase isozyme. J Biol Chem 28: 18446–18453
Mori H, Tanizawa K, Fukui T (1993) Engineered plant phosphorylases showing extraordinarily high affinity for various -glucan molecules. Protein Sci 2: 1621–1629[Medline]
Nakano K, Mori H, Fukui T (1989) Molecular-cloning of cDNA-encoding potato amyloplast alpha-glucan phosphorylase and the structure of its transit peptide. J Biochem (Tokyo) 106: 691–695
Newgard CB, Hwang PK, Fletterick RJ (1989) The family of glycogen phosphorylases: structure and function. Crit Rev Biochem Mol Biol 24: 69–99[Web of Science][Medline]
Okita TW, Greenberg E, Kuhn DN, Preiss J (1979) Subcellular localization of the starch degradative and biosynthetic enzymes of spinach leaves. Plant Physiol 64: 187–192
Olsson T, Thelander M, Ronne H (2003) A novel type of chloroplast stromal hexokinase is the major phosphorylating enzyme in the moss Physcomitrella patens. J Biol Chem 278: 44439–44447
Outlaw WH, Manchester J (1979) Guard cell starch concentration quantitatively related to stomatal aperture. Plant Physiol 64: 79–82
Preiss J (1982) Regulation of the biosynthesis and degradation of starch. Annu Rev Plant Physiol 33: 455–479[CrossRef]
Quick WP, Scheibe R, Neuhaus HE (1995) Induction of hexose-phosphate translocator activity in spinach chloroplasts. Plant Physiol 109: 113–121[Abstract]
Shimomura S, Nagai M, Fukui T (1982) Comparative glucan specificities of two types of spinach leaf phosphorylase. J Biochem (Tokyo) 91: 703–717
Smirnoff N (1993) Tansley Review 52. The role of active oxygen in the response of plants to water-deficit and desiccation. New Phytol 125: 27–58[CrossRef]
Sonnewald U, Basner A, Greve B, Steup M (1995) A second L-type isozyme of potato glucan phosphorylase: cloning, antisense inhibition and expression analysis. Plant Mol Biol 27: 567–576[CrossRef][Web of Science][Medline]
Steup M (1988) Starch degradation. In J Preiss, ed, Biochemistry of Plants. Vol 14. Carbohydrates. Academic Press, New York, pp 255–296
Steup M (1990) Starch degrading enzymes. Methods Plant Biochem 3: 103–128
Stitt M, Bulpin PV, ap Rees T (1978) Pathway of starch breakdown in photosynthetic tissues of Pisum sativum. Biochim Biophys Acta 544: 200–214[Medline]
Stitt M, Heldt HW (1981) Physiological rates of starch breakdown in isolated intact spinach chloroplasts. Plant Physiol 68: 755–761
Stitt M, Wirtz W, Gerhardt R, Heldt HW, Spencer C, Walker D, Foyer C (1985) A comparative study of metabolite levels in plant leaf material in the dark. Planta 166: 354–364[CrossRef][Web of Science]
Trethewey RN, Smith AM (2000) Starch metabolism in leaves. In RC Leegood, TD Sharkey, S von Caemmerer, eds, Advances in Photosynthesis. Vol 9. Photosynthesis: Physiology and Metabolism. Kluwer Academic Publishers, Dordrecht, the Netherlands, pp 205–231
Van Berkel J, Conrads-Strauch J, Steup M (1991) Glucan-phosphorylase forms in cotyledons of Pisum sativum L.: localization, developmental change, in vitro translation, and processing. Planta 185: 432–439[Web of Science]
Wiese A, Groner F, Sonnewald U, Deppner H, Lerchl J, Hebbeker U, Flügge UI, Weber A (1999) Spinach hexokinase I is located in the outer envelope membrane of plastids. FEBS Lett 461: 13–18[CrossRef][Web of Science][Medline]
Zeeman SC, ap Rees T (1999) Changes in carbohydrate metabolism and assimilate partitioning in starch-excess mutants of Arabidopsis. Plant Cell Environ 22: 1445–1453[CrossRef]
Zeeman SC, Northrop F, Smith AM, ap Rees T (1998a) A starch-accumulating mutant of Arabidopsis thaliana deficient in a chloroplastic starch-hydrolysing enzyme. Plant J 15: 357–365[CrossRef][Web of Science][Medline]
Zeeman SC, Umemoto T, Lue WL, Au-Yeung P, Martin C, Smith AM, Chen J (1998b) A mutant of Arabidopsis lacking a chloroplastic isoamylase accumulates both starch and phytoglycogen. Plant Cell 10: 1699–1711
This article has been cited by other articles:
|
|
 |
|
  N. Szydlowski, P. Ragel, S. Raynaud, M. M. Lucas, I. Roldan, M. Montero, F. J. Munoz, M. Ovecka, A. Bahaji, V. Planchot, et al. Starch Granule Initiation in Arabidopsis Requires the Presence of Either Class IV or Class III Starch Synthases PLANT CELL, August 1, 2009; 21(8): 2443 - 2457. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Grimaud, H. Rogniaux, M. G. James, A. M. Myers, and V. Planchot Proteome and phosphoproteome analysis of starch granule-associated proteins from normal maize and mutants affected in starch biosynthesis J. Exp. Bot., September 1, 2008; 59(12): 3395 - 3406. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
H. Satoh, K. Shibahara, T. Tokunaga, A. Nishi, M. Tasaki, S.-K. Hwang, T. W. Okita, N. Kaneko, N. Fujita, M. Yoshida, et al. Mutation of the Plastidial {alpha}-Glucan Phosphorylase Gene in Rice Affects the Synthesis and Structure of Starch in the Endosperm PLANT CELL, July 1, 2008; 20(7): 1833 - 1849. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. Deschamps, H. Moreau, A. Z. Worden, D. Dauvillee, and S. G. Ball Early Gene Duplication Within Chloroplastida and Its Correspondence With Relocation of Starch Metabolism to Chloroplasts Genetics, April 1, 2008; 178(4): 2373 - 2387. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Hofmann, D. Szakasits, A. Blochl, M. Sobczak, S. Daxbock-Horvath, W. Golinowski, H. Bohlmann, and F. M.W. Grundler Starch Serves as Carbohydrate Storage in Nematode-Induced Syncytia Plant Physiology, January 1, 2008; 146(1): 228 - 235. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Z. Deng, X. Zhang, W. Tang, J. A. Oses-Prieto, N. Suzuki, J. M. Gendron, H. Chen, S. Guan, R. J. Chalkley, T. K. Peterman, et al. A Proteomics Study of Brassinosteroid Response in Arabidopsis Mol. Cell. Proteomics, December 1, 2007; 6(12): 2058 - 2071. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Edner, J. Li, T. Albrecht, S. Mahlow, M. Hejazi, H. Hussain, F. Kaplan, C. Guy, S. M. Smith, M. Steup, et al. Glucan, Water Dikinase Activity Stimulates Breakdown of Starch Granules by Plastidial beta-Amylases Plant Physiology, September 1, 2007; 145(1): 17 - 28. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. L. Nielsen, A. L. Hogh, and J. Emmersen DeepSAGE--digital transcriptomics with high sensitivity, simple experimental protocol and multiplexing of samples Nucleic Acids Res., November 14, 2006; 34(19): e133 - e133. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Lu, J. M. Steichen, J. Yao, and T. D. Sharkey The Role of Cytosolic {alpha}-Glucan Phosphorylase in Maltose Metabolism and the Comparison of Amylomaltase in Arabidopsis and Escherichia coli Plant Physiology, November 1, 2006; 142(3): 878 - 889. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. Alonso-Casajus, D. Dauvillee, A. M. Viale, F. J. Munoz, E. Baroja-Fernandez, M. T. Moran-Zorzano, G. Eydallin, S. Ball, and J. Pozueta-Romero Glycogen Phosphorylase, the Product of the glgP Gene, Catalyzes Glycogen Breakdown by Removing Glucose Units from the Nonreducing Ends in Escherichia coli. J. Bacteriol., July 1, 2006; 188(14): 5266 - 5272. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. E. Weise, S. M. Schrader, K. R. Kleinbeck, and T. D. Sharkey Carbon Balance and Circadian Regulation of Hydrolytic and Phosphorolytic Breakdown of Transitory Starch Plant Physiology, July 1, 2006; 141(3): 879 - 886. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Balmer, W. H. Vensel, N. Cai, W. Manieri, P. Schurmann, W. J. Hurkman, and B. B. Buchanan A complete ferredoxin/thioredoxin system regulates fundamental processes in amyloplasts PNAS, February 21, 2006; 103(8): 2988 - 2993. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Borland, S. Elliott, S. Patterson, T. Taybi, J. Cushman, B. Pater, and J. Barnes Are the metabolic components of crassulacean acid metabolism up-regulated in response to an increase in oxidative burden? J. Exp. Bot., January 1, 2006; 57(2): 319 - 328. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T.-S. Yu, S. C. Zeeman, D. Thorneycroft, D. C. Fulton, H. Dunstan, W.-L. Lue, B. Hegemann, S.-Y. Tung, T. Umemoto, A. Chapple, et al. {alpha}-Amylase Is Not Required for Breakdown of Transitory Starch in Arabidopsis Leaves J. Biol. Chem., March 18, 2005; 280(11): 9773 - 9779. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. M. Smith, D. C. Fulton, T. Chia, D. Thorneycroft, A. Chapple, H. Dunstan, C. Hylton, S. C. Zeeman, and A. M. Smith Diurnal Changes in the Transcriptome Encoding Enzymes of Starch Metabolism Provide Evidence for Both Transcriptional and Posttranscriptional Regulation of Starch Metabolism in Arabidopsis Leaves Plant Physiology, September 1, 2004; 136(1): 2687 - 2699. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| ASPB Publications | PLANT PHYSIOLOGY® | THE PLANT CELL | |
|---|---|---|---|
-Glucan Phosphorylase Is Not Required for Starch Degradation in Arabidopsis Leaves But Has a Role in the Tolerance of Abiotic Stress
