| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
|
First published online September 19, 2008; 10.1104/pp.108.127969 Plant Physiology 148:1614-1629 (2008) © 2008 American Society of Plant Biologists Alterations in Cytosolic Glucose-Phosphate Metabolism Affect Structural Features and Biochemical Properties of Starch-Related Heteroglycans1,[W]Max-Planck Institute of Molecular Plant Physiology (A.N.-N., A.R.F.) and Department of Plant Physiology (J.F., J.A., M.S., M.S.), University of Potsdam, 14476 Potsdam-Golm, Germany
The cytosolic pools of glucose-1-phosphate (Glc-1-P) and glucose-6-phosphate are essential intermediates in several biosynthetic paths, including the formation of sucrose and cell wall constituents, and they are also linked to the cytosolic starch-related heteroglycans. In this work, structural features and biochemical properties of starch-related heteroglycans were analyzed as affected by the cytosolic glucose monophosphate metabolism using both source and sink organs from wild-type and various transgenic potato (Solanum tuberosum) plants. In leaves, increased levels of the cytosolic phosphoglucomutase (cPGM) did affect the cytosolic heteroglycans, as both the glucosyl content and the size distribution were diminished. By contrast, underexpression of cPGM resulted in an unchanged size distribution and an unaltered or even increased glucosyl content of the heteroglycans. Heteroglycans prepared from potato tubers were found to be similar to those from leaves but were not significantly affected by the level of cPGM activity. However, external glucose or Glc-1-P exerted entirely different effects on the cytosolic heteroglycans when added to tuber discs. Glucose was directed mainly toward starch and cell wall material, but incorporation into the constituents of the cytosolic heteroglycans was very low and roughly reflected the relative monomeric abundance. By contrast, Glc-1-P was selectively taken up by the tuber discs and resulted in a fast increase in the glucosyl content of the heteroglycans that quantitatively reflected the level of the cytosolic phosphorylase activity. Based on 14C labeling experiments, we propose that in the cytosol, glucose and Glc-1-P are metabolized by largely separated paths.
During photosynthesis, most plant species utilize a major proportion of the carbon fixed to accumulate transitory starch, which is deposited in the stromal space of the chloroplasts. Chloroplastic starch is degraded in the subsequent dark period and, thereby, plants are able to sustain growth and developmental processes in the absence of photosynthetic carbon assimilation (Smith et al., 2005  ; Fettke et al., 2007; Smith and Stitt, 2007; Zeeman et al., 2007a, 2007b). Starch mobilization appears to be initiated by a combined action of starch phosphorylating dikinases (Ritte et al., 2002; Hejazi et al., 2008) and various hydrolases, such as plastidial β-amylase isozymes (Scheidig et al., 2002; Fulton et al., 2008) and a debranching isoenzyme (ISA3; Edner et al., 2007). This process leads to the formation of maltose and a series of maltodextrins, which are further metabolized by the plastidial disproportionating isozyme 1 (DPE1) to yield Glc (Chritchley et al., 2001). However, most of the starch-derived neutral sugars are exported by the chloroplasts as maltose (Weise et al., 2004). Following the transport into the cytosol via selective transporters (designated MEX and pGT; Weber et al., 2000; Niittylä et al., 2004), both Glc and maltose are converted to phosphorylated sugars, which fuel a variety of biosynthetic processes, including the formation of Suc and cell wall constituents (Reiter, 2008). The initial reactions of this path are unexpectedly complex and contain, as an indispensable step, a reversible glucosyl transfer from maltose to a glycan-type acceptor and the release of the other maltose-derived glucosyl residue (the one containing the reducing end of the former disaccharide) as Glc. This reaction is catalyzed by a cytosolic transglucosidase (EC 2.4.1.25; also designated amylomaltase, glucosyl transferase, or DPE2). DPE2 is a modular protein containing two putative N-terminal carbohydrate-binding modules and a C-terminal domain similar to that of glycoside hydrolases of family 77, which, however, is split by a large insertion (Steichen et al., 2008). Arabidopsis (Arabidopsis thaliana) mutants deficient in DPE2 differ from wild-type plants by elevated levels of both maltose and starch and by a strongly reduced growth (Chia et al., 2004; Lu and Sharkey, 2004). Based on sequence analyses, both DPE2 and MEX are widely distributed in higher and lower plants, including Ostreococcus tauri and Chlamydomonas reinhardtii (Deschamps et al., 2008; Steichen et al., 2008). This implies that both proteins exert an important function in starch metabolism throughout the plant kingdom.
Under in vitro conditions, DPE2 catalyzes the repetitive transfer of glucosyl residues from maltose molecules to nonreducing ends of glycogen (Chia et al., 2004
On the basis of several independent lines of evidence, subfraction I is likely to be the in vivo substrate of DPE2 in leaves and, therefore, may be intimately involved in the cytosolic metabolism of starch-derived monosaccharides and disaccharides. Recombinant DPE2 transfers glucosyl residues from maltose to subfraction I and from subfraction I to Glc, thereby yielding Glc and maltose, respectively. By contrast, DPE2 does not utilize subfraction II as glucosyl acceptor or as glucosyl donor (Fettke et al., 2006
According to this path (Fig. 1), the cytosolic pools of both maltose and Glc are immediately or via a short sequence of reactions linked to the heteroglycans. DPE2 transfers a glucosyl residue from maltose to the heteroglycans and also from the glycans to Glc, yielding maltose. When using maltose as glucosyl donor, the glucosyl moiety containing the former reducing end of the disaccharide is released as Glc, which enters the cytosolic pool as does Glc generated from starch inside the chloroplast, and is subsequently exported to the cytosol. By the action of hexokinase, the cytosolic Glc pool is phosphorylated to yield Glc-6-P, which, subsequently, is converted to Glc-1-P by the cytosolic isozyme(s) of phosphoglucomutase (PGM; EC 2.7.5.1). The two Glc-Ps enter many pathways, among which, in quantitative terms, the biosynthesis of Suc and the formation of cell wall constituents are most prominent (Scheible and Pauly, 2004 ). Alternatively, Glc-1-P can act as glucosyl donor for the Pho2-mediated transfer of glucosyl residues to the heteroglycans. Due to the reversibility of the Pho2-catalyzed reaction, the enzyme can also utilize the heteroglycan as a glucosyl donor and orthophosphate as an acceptor, catalyzing the formation of Glc-1-P.
The cyclic process outlined in Figure 1, which includes the interconversion of Glc-6-P and Glc-1-P, enables the cell to balance varying influxes of monosaccharides and disaccharides from the numerous chloroplasts into the cytosol. Recently, several transgenic lines of potato were phenotypically characterized that possess altered cytosolic phosphoglucomutase (cPGM) activity. Some of these lines express a PGM from Escherichia coli in addition to the endogenous PGM isoforms. Expression of the transgenes was controlled either by the tuber-specific B33 patatin promoter or the tissue-constitutive 35S promoter. In leaves possessing an elevated cPGM activity, photosynthesis rates were essentially unchanged but the Suc content was increased. Interestingly, the level of maltose (and that of isomaltose as well) was significantly decreased in all transgenic lines. In addition, the content of minor monosaccharides, such as Gal and Ara, was lowered (Lytovchenko et al., 2005a
The reason for analyzing heteroglycans from potato tuber was 2-fold. First, almost nothing is known about the structure and functions of these glycans in heterotrophic tissues. In this system, major carbon fluxes are directed from the cytosol toward the plastid, whereas in autotrophic cells, fluxes from the chloroplast into the cytosol are dominant. Second, exogeneous sugars are efficiently taken up by tuber discs and are utilized for a massive reserve starch biosynthesis inside the amyloplast (Geiger et al., 1998
PGM Activity in the Transgenic Potato Plants
Four previously generated transgenic potato lines that either overexpress or underexpress the cPGM were grown under controlled conditions together with wild-type plants. Source leaves or growing tubers freshly removed from the mother plants were harvested, and the total extractable PGM activities were monitored (Table I
). In leaves, the protein-based total PGM activity of the most efficiently overexpressing line (denoted line B here) and the most strongly reduced line (denoted line C here) differ by approximately 5-fold (the original designations of these lines are provided in "Materials and Methods"). For the activity of the cPGM isozyme(s), this range is likely to be even larger, as it represents approximately 55% of the total extractable enzyme activity (Tauberger et al., 2000
Next, we determined the PGM isoform patterns of leaves (Fig. 2A ) and tubers (Fig. 2B) by native PAGE followed by an enzyme activity staining that is based on the PGM-dependent conversion of Glc-1-P to Glc-6-P and two subsequent redox reactions that finally result in the formation of insoluble formazan (Kofler et al., 2000 ). In all samples, three bands of PGM activity were resolved, all of which strictly depend upon the addition of Glc-1-P (data not shown). In the transgenic lines that express the PGM from E. coli in the cytosol (lines A and B; Lytovchenko et al., 2005b), the transgenic PGM comigrated with the most slowly moving endogenous PGM form (band 3; Fig. 2, A and B). A strong overexpression of the bacterial PGM tends to diminish the resolution of the three bands of activity. In antisense lines (C and D), the staining of the slowest moving band is diminished (Fig. 2, A and B). These results suggest that band 3 represents the cPGM isozyme. However, due to the instability of the PGM activity from potato leaves or tubers, verification of this assumption by nonaqueous fractionation was unsuccessful.
Therefore, we compared the PGM isozyme pattern from potato with that of leaves from Arabidopsis plants, and we included a mutant that is deficient in the plastidial isoform (Caspar et al., 1985 ). For leaves from Arabidopsis wild-type plants, three bands of PGM activity were observed that, according to the order of decreasing mobilities, were designated bands I to III. In the mutant, the two slower moving bands (bands II and III; i.e. the cytosolic forms) were present but the fast moving band (band I; the plastidial isozyme) was undetectable (Fig. 2C). Interestingly, essentially the same PGM isozyme pattern was observed when leaf extracts were separated by a continuous electrophoresis in starch gels (Caspar et al., 1985). In summary, it is highly likely that in extracts from both leaves and tubers from potato, band 3 is located in the cytosol whereas band 1 is plastidial. The subcellular location of band 2 remains to be defined. This isoform could be attributed to the cytosol (like band II from Arabidopsis) or it could represent a modified plastidial isozyme.
Some features of the photometrically assayed PGM activity, as shown in Table I and the zymograms (Fig. 2), should be noted. In the photometric assay, the total PGM activity is monitored under steady-state conditions using a saturating substrate (i.e. Glc-1-P) concentration. All PGM isoforms contribute to the measured total enzyme activity, but it is unknown whether the assay conditions used are optimal for all isoforms. Therefore, it is not known whether the total PGM activity measured equals the sum of the Vmax values of all PGM isoforms. For the zymograms, equal amounts of buffer-soluble proteins were applied to each lane. PGM activity detection was optimized to obtain the most sensitive enzyme activity staining, the highest possible resolution of the PGM isoforms, and a low background staining as well. Based on these criteria, the native PAGE system originally described by Davis (1964) We also tested both potato leaf and tuber extracts from the four transgenic lines and from wild-type plants for possible variation in transglucosidase and phosphorylase isozyme patterns; however, neither qualitative nor quantitative differences were observed among all the lines. Therefore, we conclude that the altered expression of the cPGM does not noticeably affect the activities of the two subfraction I-related glucosyl transferases, Pho2 and DPE2 (Supplemental Fig. S1).
Heteroglycans were isolated from the leaves of two overexpressing lines (lines A and B), the two underexpressing lines (lines C and D), and the wild type. Leaves were harvested both in the middle of the light period (8 h after the onset of the illumination) and in the middle of the dark period (4 h after the beginning of the dark period). Subfractions I and II were separated by FFF and subjected to acid hydrolysis. The monomeric patterns of the hydrolysates were determined by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD; Fig. 3
). In subfraction I, Gal and Ara were the two most prominent compounds, whereas Fuc, Rha, Glc, Xyl, and Man were minor constituents (compare with Fettke et al., 2005b
As shown previously, both subfractions I and II possess a relatively wide and overlapping size distribution (Fettke et al., 2005a , 2005b). In wild-type potato leaves, the size distribution of subfraction I increases during the dark period, whereas that of subfraction II remains constant during the light-dark period (Fettke et al., 2005b). For a more detailed analysis, SHGL was prepared from leaves of the four transgenic lines harvested at the middle of the light period and equal amounts (80 µg of Glc equivalents each) were subjected to FFF, and both the multi-laser light scattering and the delta refractive index (MALLS-DRI) were monitored. As controls, wild-type leaves were harvested and processed under identical conditions (Fig. 4 ). Two observations were made. First, in SHGL fractions derived from the two overexpressing lines (lines A and B), the size distribution of subfraction I was shifted toward the lower molecular mass region (Fig. 4A). Second, in these lines, the ratio between subfractions I and II was decreased. By contrast, both subfractions I and II from underexpressing lines (lines C and D) were quantitatively and qualitatively essentially undiscernible from those of the wild type. The same differences between SHGL preparations derived from line A or B and from the wild-type control were observed when the glycans were prepared from leaves that had been harvested during darkness. Similarly, the two underexpressing lines (lines C and D) did not differ from the wild type (data not shown).
Taken together, the data presented in Figures 3 and 4 are fully consistent with the cyclic path outlined in Figure 1: elevated levels of cPGM activity favor the phosphorylase-mediated conversion of glucosyl residues from subfraction I into the cytosolic hexose-P pools. However, this increased flux from the cytosolic heteroglycans is not restricted to the glucosyl residues; rather, it is accompanied by a more general decrease in the total amount and size of the cytosolic heteroglycans. By contrast, more glucosyl residues are retained in subfraction I when the levels of cPGM activity are decreased. Under these conditions, neither the amount nor the size distribution of subfraction I is altered (Fig. 4B).
The monomer patterns of the SHGs were also analyzed (Supplemental Fig. S2, A and B). In these glycans, Glc is the dominant monomer, but it is more prominent during the light period than during darkness. This effect was also observed in the SHGS preparations from all four transgenic lines. In comparison with the wild type, the relative Glc levels of the mutant-derived SHGs preparations were either unchanged or lower. As SHGS comprises several glycan pools that reside in different compartments, however, these differences are difficult to explain at the molecular level (Fettke et al., 2005a
Growing potato tubers are characterized by a massive starch biosynthetic flux and act as major sinks of the entire plant (Geigenberger et al., 2004 When using essentially the same procedure that has been established for leaves from Arabidopsis or potato, the SHGL preparation obtained still contained minor protein bands. As revealed by tryptic digestion and matrix-assisted laser-desorption ionization mass spectrometry analyses, these proteins were unrelated to any glycoproteins but rather represented major storage proteins of the tuber that had been incompletely removed during deproteinization of the glycans (Supplemental Fig. S3). By contrast, noticeable protein contaminations were never observed in heteroglycans isolated from any leaf material.
As contaminating proteins of the tuber-derived SHGL preparations could interfere both with FFF-MALLS analyses and with the quantification of the total monosaccharide content, several procedures were tested in an attempt to achieve complete removal of contaminating proteins. Treatment of SHGL with proteinase K was found to be most efficient (Supplemental Fig. S3). Presumably, the autodegradation catalyzed by proteinase K is advantageous for the enzymatic deproteinization of the tuber-derived SHGL (Bajorath et al., 1988
Following proteinase K treatment, SHGL preparations from growing tubers of the various transgenic lines (and from wild-type tubers as well) were separated into subfractions I and II either by FFF or by reaction with the β-glucosyl Yariv reagent. As revealed by HPAEC-PAD analyses, subfractions I and II isolated from wild-type tubers possess similar monomer patterns as the respective subfractions from wild-type leaves (see below). Likewise, the ratio and size distribution of subfractions I and II, as revealed by FFF-MALLS-DRI, were essentially as described previously for leaves (data not shown; Fettke et al., 2005b
Given that it is likely that only major structural alterations of the heteroglycans are reflected by changes in the size distribution or the monomer pattern, we next attempted a more subtle analysis based on evaluating the metabolic fate of the supplied radiolabel. Labeling experiments were performed with tuber discs from wild-type plants and both a cPGM-overexpressing line (line A) and a cPGM-underexpressing transgenic line (line C). All discs were prepared from growing potato tubers that were removed from the mother plant immediately before the experiment. Discs were incubated with uniformally 14C-labeled Glc in sealed flasks, and the CO2 released was trapped in an alkaline solution. At intervals, the alkaline solution was substituted and discs were removed. Subsequently, they were analyzed to determine the distribution of label. The data obtained are compiled in Table II
. Total uptake of 14C-labeled Glc continued over the entire incubation period, the rate of uptake being approximately the same in line A as in the wild type but approximately 20% lower in the antisense line C. Interestingly, in keeping with the changes in steady-state metabolite levels previously recorded in this antisense lines (Fernie et al., 2002b
Next, we evaluated the 14C incorporation into heteroglycans within the exact same tuber material used for the experiments described above. SHGL was separated into subfractions I and II by treatment with the β-glucosyl Yariv reagent (Yariv et al., 1962 ). As demonstrated previously, the apoplastic subfraction II strongly interacts with this reagent, whereas the cytosolic subfraction I is nonreactive (Fettke et al., 2004). As incorporation of radiolabel was low, analyses were restricted to the longer incubation periods of the experiment described above. Incorporation into subfraction I was greater than that into subfraction II. However, no significant differences were observed between wild-type controls and the two transgenic lines. The antisense line (line C) exhibited a slightly decreased labeling of subfraction I in comparison with the wild-type control (Fig. 5A
).
We then determined the labeling pattern of the individual constituents of SHGL (Fig. 5B). Given the low rate of 14C incorporation into the heteroglycan, analyses were restricted to the 240-min incubation period. For this purpose, we performed an acid hydrolysis, and equal amounts of the monosaccharides released were applied to an HPAEC column. Eluate fractions were collected that each represented a distinct monosaccharide, and the radiolabel content of each fraction was monitored. Galactose, which is the most prominent constituent of SHGL, contained the majority of the 14C label, whereas incorporation into the other monosaccharides was minor. In the antisense line (line C) and in the overexpressing line (line A), labeling of the various monosaccharides did not differ noticeably from that of the wild type. In summary, the 14C incorporation into the various monosaccharides roughly reflects their relative abundance within the heteroglycans. This strongly suggests that even during the prolonged incubation, only a small proportion of the labeled Glc is partitioned to the various constituents of the glycans, without any clear preference being noticeable. When comparing the labeling of the heteroglycans with the 14C incorporation into major cellular constituents of the tuber, it becomes apparent that, at least under the conditions used here, the Glc-driven heteroglycan biosynthesis represents only a very minor intracellular flux within the potato tuber (their total 14C content per gram fresh weight is less than 1% of that found in starch). This low labeling of the heteroglycans also has an important methodological implication: it clearly indicates that the heteroglycans isolated from potato tubers lack, to any noticeable extent, starch-derived contaminations; otherwise, a much higher labeling, especially of the glucosyl residues within the heteroglycans, would be expected.
Potato tuber discs were incubated with Glc-1-P (or, alternatively, with equimolar concentrations of other sugars) and, subsequently, the monomer pattern of SHGL was analyzed by HPAEC-PAD (Fig. 6A ). Following a 60-min incubation with Glc-1-P, the glucosyl content of SHGL was increased significantly. This effect was not detectable when Glc-1-P was replaced by Glc-6-P, Glc, or Suc. In a further control experiment, discs were incubated with a mixture containing Glc, ATP, and MgCl2 in order to facilitate the in situ formation of Glc-6-P by any endogenous extracellular hexokinase. However, the effect of external Glc-1-P could not be mimicked (Fig. 6A). In summary, the data shown in Figure 6A suggest that external Glc-1-P and free Glc exert different effects on the cytosolic carbon fluxes.
The above conclusion was supported by further experiments. Discs from wild-type tubers were incubated for varying periods of time with unlabeled Glc-1-P. As a control, Glc-1-P was omitted. For each incubation time, SHGL preparations were isolated and their monosaccharide patterns were analyzed by HPAEC-PAD. The Glc-1-P-dependent increase in the glucosyl content of SHGL is remarkably fast, as most of the change in the monosaccharide pattern occurs during 30 min following the onset of the incubation (Fig. 6B). Subsequently, the increase in the glucosyl content slows, and following prolonged incubation with Glc-1-P (for 2 or 3 h) the glucosyl content even decreases (data not shown). As revealed by native PAGE, the phosphorylase and the DPE2 patterns were not significantly altered during the incubation with any of the compounds supplied (Supplemental Fig. S4).
According to the path proposed for mesophyll cells (Fig. 1), the cytosolic phosphorylase isozyme (Pho2) catalyzes the reversible glucosyl transfer from Glc-1-P to the heteroglycans. If this path is also functional in tuber cells, the Glc-1-P-dependent increase in the glucosyl content of SHGL should be correlated with the level of Pho2 activity. In order to test this assumption, tuber discs from four transgenic potato lines that possess elevated (lines 3 and 4) or reduced (lines 1 and 2) levels of the cytosolic phosphorylase were incubated with Glc-1-P. In a previous publication, these lines were characterized in detail (Fettke et al., 2005b
In order to further substantiate this conclusion, two additional experiments were performed. In the first experiment, wild-type tuber discs were incubated in three mixtures containing equimolar concentrations of Glc-1-P or Glc or lacking any carbohydrate. SHGL was isolated from each set of discs and was then resolved into subfractions I and II by treatment with β-glucosyl Yariv reagent. The increase in the glucosyl content was restricted to the Yariv nonreactive subfraction (i.e. the cytosolic subfraction I), and this increase was only observed following incubation with Glc-1-P (Fig. 7B). In the second experiment, SHGL was treated with an endo-  -1,5-arabinanase from Aspergillus niger and, subsequently, the residual (>10 kD) heteroglycan was freed of low molecular mass glycans (<10 kD) and was then subjected to acid hydrolysis. The treatment with endo-arabinanase that selectively cleaves inter-Ara linkages has previously been applied successfully to characterize cytosolic heteroglycans from leaves of both Arabidopsis and potato (Fettke et al., 2005b, 2006). The monosaccharide pattern of the residual SHGL is shown in Figure 7C. Two control experiments were performed. In the first control, SHGL was treated in exactly the same way except that the endo-arabinanase was omitted (Fig. 7C). In the second control, SHGL was replaced by an equal amount of glycogen. The incubation with the arabinanase did not result in any noticeable hydrolytic activity (data not shown). Due to the action of the arabinanase, the >10-kD heteroglycan contains less Ara, Rha, Gal, and Glc. However, the loss of Glc content exceeds that of the other monosaccharides. Similar results have been obtained for SHGL that had been isolated from freshly harvested potato leaves (Fettke et al., 2005b). Since phosphorylases catalyze the reversible transfer of glucosyl residues from Glc-1-P to the nonreducing ends of acceptor molecules, it is reasonable to assume that most of the glucosyl residues observed in subfraction I are placed at the periphery of the polyglycan molecules. The susceptibility of these glucosyl residues (and of a few other monosaccharide residues as well) to arabinanase treatment clearly indicates that they have been transferred to a heteroglycan rather than to an as yet unidentified homoglucan.
The data shown in Figures 6 and 7 clearly demonstrate that external Glc and Glc-1-P differently affect the monomer pattern of cytosolic heteroglycans. The increase in the glucosyl content of subfraction I is strictly dependent on Glc-1-P and is mediated by the cytosolic Pho2 (Fig. 7). Therefore, it is likely that following uptake, the external Glc-1-P directly enters the cytosolic pool of this sugar phosphate.
The import of Glc-1-P into intact amyloplasts isolated from tubers or suspension-cultured cells from potato has been reported (Kosegarten and Mengel, 1994
In this article, we have analyzed cytosolic heteroglycan-related processes in two distinct systems of potato. One system is the fully developed leaf, which acts as the major source organ of the entire plant. Within this system, carbon fixed during photosynthesis is exported from the chloroplast to the cytosol utilizing either intermediates of the Calvin cycle (during the light period) or products formed by the degradation of transitory starch (during the dark period). Presumably, at least two paths exist by which plastidial metabolites are linked to the cytosolic heteroglycans: one predominantly acts during net starch degradation and includes both maltose and Glc transporters, and the other one is mediated by the triose-P transporter and links Calvin cycle intermediates with the cytosolic heteroglycans. The latter process is consistent with the fact that cytosolic heteroglycans have been isolated from starch-deficient Arabidopsis mutants (Fettke et al., 2005a ) and, therefore, that the in vivo biosynthesis of the glycans proceeds even if starch-derived monosaccharides or disaccharides are not available. Furthermore, in vivo labeling of Arabidopsis wild-type plants using 14CO2 clearly indicated that the labeled carbon is incorporated into the glycans during the light period (Fettke et al., 2005a). The evidence for the DPE2-mediated involvement of the heteroglycans in the cytosolic maltose metabolism is based on both in vitro and in vivo studies and includes the massive structural alterations observed in heteroglycans from DPE2-deficient Arabidopsis mutants (Fettke et al., 2006).
In the leaf system, heteroglycans were isolated from transgenic plants that possess altered levels of the cPGM activity. The total extractable PGM activity was resolved into three isoforms, and the same pattern was obtained for tubers (Fig. 2, A and B). The fastest moving isoform appears to be plastidial, whereas the slowest moving form is attributed to the cytosol. The subcellular location of the third band (the one having an intermediate mobility) remains uncertain. A similar PGM pattern was obtained for Arabidopsis wild-type leaves. In the mutant lacking the plastidial PGM isozyme, the fastest moving form was undetectable (Fig. 2C). For Arabidopsis, two genes encoding cPGM isozymes (At1g23190 and At1g70730) have been identified. Single knockout mutants in which either At1g23190 or At1g70730 is nonfunctional possess an approximately 50% effect on starch and Suc content (J. Fettke, unpublished data). Therefore, the two gene products appear to exert redundant rather than selective metabolic functions. In transgenic potato lines that express the E. coli-derived PGM in the cytosol (overexpressing lines A and B), the transgenic product comigrates with the slowest moving endogenous PGM isoform (Fig. 2, A and B). In the two antisense lines, the slowest moving cPGM isoform is most efficiently repressed (Fig. 2). It is important to note that, in contrast to Arabidopsis, the potato genome appears to encode only two PGMs. In keeping with this partial purification, studies revealed two major distinct peaks (Takamiya and Fukui, 1978
Both elevated and lowered levels of cPGM activity affect the cytosolic heteroglycans (i.e. subfraction I), whereas the apoplastic water-soluble glycans (subfraction II) remain unchanged (Figs. 3 and 4). This result is consistent with the scheme presented in Figure 1, as subfraction I is linked, via a series of reversible reactions, with the cytosolic pools of Glc-6-P and Glc-1-P. Elevated levels of PGM activity (lines A and B) favor the interconversion of the two Glc monophosphates and result in an increased flux into Suc (Lytovchenko et al., 2005a
The other system used in this study is growing tubers, which constitute the most active sinks of potato plants. In tubers, parenchyma cells import source-derived sugars and direct most of the reduced carbon toward the amyloplasts to support a massive starch biosynthetic flux (Geigenberger et al., 2004
The supply of Glc-1-P results in totally different carbon fluxes from those following the supply of Glc. Glc-1-P seems to be selectively taken up (Fig. 8) and then immediately enriches the cytosolic Glc-1-P pool, providing the cytosolic phosphorylase (Pho2) isozyme with an increased substrate supply. Given the reversibility of the Pho2-catalyzed glucosyl transfer, the label in Glc residues of subfraction I increases rapidly and massively (Fig. 6). This increase correlates, quantitatively, with the level of the cytosolic phosphorylase activity, as indicated by the data obtained with Pho2-overexpressing or -underexpressing lines (Fig. 7A). As would be anticipated from Figure 1, these effects are restricted to subfraction I, as the cytosolic phosphorylase does not possess immediate access to the apoplastic glycans. The conclusions drawn from the data shown in Figures 6 to 8
Our results indicate that Glc-1-P is taken up by an alternative route to Glc and that the differences observed in our experiments reflect the differential availability of these compounds within the cytosol. Intriguingly, however, despite this apparent separation, the Glc-1-P-dependent pathway remains capable of supporting a massive carbon flux into starch (Fig. 8). Interestingly, transgenic potato plants that contain low activities of both the cPGM and plastidial PGM unexpectedly exhibit a phenotype that is similar to that of wild-type plants (Fernie et al., 2002a ). It is not inconceivable that under these conditions an intracellular carbon flux is dominant that resembles the Glc-1-P-dependent flux, as observed in this study. At present, experiments are being performed in our laboratories that should allow this hypothesis to be tested.
Plant Materials
Potato plants (Solanum tuberosum ‘Desiree’) and Pho2 transgenic lines were grown under controlled conditions as described (Fettke et al., 2005b
Leaf material was frozen in liquid nitrogen and broken using a mortar. All subsequent steps were performed at 4°C. Per 1 g fresh weight, 1 mL of extraction buffer (100 mM HEPES-NaOH, pH 7.5, 1 mM EDTA, 2 mM dithioerythritol, 10% [v/v] glycerol, and 0.5 mM phenylmethylsulfonyl fluoride) was added to the powdered leaf material, and the resulting homogenate was centrifuged for 12 min at 20,000g. Tuber material (1–3 g fresh weight each) was homogenized in 5 mL of extraction buffer using an Ultra Turrax homogenizer. Following centrifugation (as above), the supernatant was passed through a nylon net (100-µm mesh) and the filtrate (designated the crude extract) was used for enzyme activity assays, protein quantification, and native PAGE. Protein was quantified using the microassay of Bradford (1976)
For native PAGE (7.5% [w/v] total monomer concentration), the Mini-Protean II apparatus (Bio-Rad) was used. Electrophoresis was performed at 4°C using a precooled electrode buffer that was continuously stirred during electrophoresis. PGM activity was resolved using the native discontinuous PAGE system originally described by Davis (1964)
Leaves were quickly frozen in liquid nitrogen. Aliquots of the frozen leaf material (2 g each) were broken in a mortar, and the SHG was extracted in 10 mL of 20% (v/v) ethanol. The entire isolation and fractionation procedure is described elsewhere (Fettke et al., 2005a
For photometric determination of the PGM, an assay similar to that of the phosphorylase (Steup and Latzko, 1979
Developing tubers were removed from 10-week-old plants. A longitudinal core (10 mm diameter) was taken, and the core was sliced into discs (1 mm thickness) that were washed three times in precooled buffer (10 mM MES-KOH, pH 6.5). Subsequently, the tuber discs (20 each) were placed into a 100-mL Erlenmeyer flask containing 10 mL of incubation medium consisting of 50 mM unlabeled Glc and 10 µCi [U-14C]Glc (specific radioactivity, 0.74 MBq mmol–1) in 10 mM MES-KOH, pH 6.5. The flasks were sealed with Parafilm, and the CO2 released was captured in a KOH trap for quantification by liquid scintillation counting. Incubation of the tuber discs was terminated by washing with cold water. Subsequently, discs were frozen in liquid nitrogen and were stored at –80°C until sample processing. Metabolites were extracted by a series of incubation steps (10 min each). The discs were extracted using a series of boiling solvents (2 mL each) consisting of 80% (v/v) ethanol, 50% (v/v) ethanol, 20% (v/v) ethanol, water, and, finally, 80% (v/v) ethanol. The supernatants were combined and dried under vacuum. Subsequently, the pellet was rapidly resuspended in water (2 mL) and an aliquot was used to monitor radioactivity by liquid scintillation counting. Hexoses and Suc from the ethanol-soluble fraction and starch, protein, and cell wall matter were separated by enzymatic degradation as described by Carrari et al. (2006)
Discs (8 mm diameter, approximately 2 mm thickness) were prepared from freshly harvested tubers and were incubated at room temperature in one of the following mixtures (10 discs each in 10 mL), all containing 10 mM citrate-NaOH, pH 6.5, as buffer: (1) buffer; (2) 25 mM Glc; (3) 25 mM Glc-1-P; (4) 25 mM Glc-6-P; (5) 25 mM Glc, 25 mM ATP, and 2.5 mM MgCl2; (6) 25 mM Suc. At intervals (0, 30, and 60 min), five discs each were withdrawn and frozen in liquid nitrogen. SHG were isolated as described above. Buffer-soluble proteins were extracted from five discs each and were subjected to native PAGE. Gels were stained for phosphorylase or DPE2 activity as described above.
Using exactly the same experimental conditions as described above, tuber discs were incubated with 50 mM [U-14C]Glc, [U-14C]Suc, or 10 mM [U-14C]Glc-1-P in 50 mM citrate-NaOH, pH 6.5. In addition, 10 mM [U-14C]Glc-1-P and unlabeled 5 mM Glc-6-P and Gal-1-P, respectively were mixed. To each mixture, 74 kBq of radioactivity was included. After incubation at room temperature for 45 and 60 min, four discs each were withdrawn, washed with water, and frozen in liquid nitrogen. For sample processing, four discs were homogenized in 5 mL of precooled 20% (v/v) ethanol using an Ultra Turrax homogenizer. Subsequently, the samples were centrifuged (10 min at 10,000g, 4°C) and the supernatant was collected separately. The resulting pellets were washed with 1 mL of water and centrifuged as above. The 14C label in the pellet, representing predominantly starch, was monitored using a liquid scintillation counter. The supernatants were combined and 100% (v/v) ethanol was added to give a final concentration of 70% (v/v). Following incubation overnight at –20°C, the samples were centrifuged (as above), and in both the resulting pellet (designated the polysaccharides, essentially SHGL) and the supernatant (containing monosaccharides and oligosaccharides, SHGS, and other constituents soluble in 70% [v/v] ethanol), the 14C-label was monitored using a liquid scintillation counter.
The following materials are available in the online version of this article.
This work was supported by the Interdisciplinary Center for Advanced Protein Technologies of the University of Potsdam. Received August 11, 2008; accepted September 8, 2008; published September 19, 2008.
1 This work was supported by the Deutsche Forschungsgemeinschaft: Sonderforschungsbereich 429 "Molecular Physiology, Energetics, and Regulation of Primary Metabolism in Plants," Teilprojekte A11 (A.R.F.), and B2 (J.F. and M. Steup). 
2 Present address: Department of Biochemistry, Faculty of Agriculture and Biology, Warsaw University of Life Sciences, 02–776 Warszawa, Poland. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Martin Steup (msteup{at}rz.uni-potsdam.de).
[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.108.127969 * Corresponding author; e-mail msteup{at}rz.uni-potsdam.de.
Appeldoorn NJG, de Bruijn SM, Koot-Gronsveld EAM, Visser RGF, Vreugdenhil D, van der Plas LHW (1997) Developmental changes of enzymes involved in conversion of sucrose to hexose-phosphate during early tuberisation of potato. Planta 202: 220–226[CrossRef][Web of Science] Bajorath J, Hinrichs W, Saenger W (1988) The enzymatic activity of proteinase K is controlled by calcium. Eur J Biochem 176: 441–447[Web of Science][Medline] Benfey PN, Chua NH (1990) The cauliflower mosaic virus 35S promoter: combinatorial regulation of transcription in plants. Science 16: 959–966 Bradford MM (1976) A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248–254[CrossRef][Web of Science][Medline] Carrari F, Baxter C, Usadel B, Urbanczyk-Wochniak E, Zanor MI, Nunes-Nesi A, Nikiforova V, Centero D, Ratzka A, Pauly M, et al (2006) Integrated analysis of metabolite and transcript levels reveals the metabolic shifts that underlie tomato fruit development and highlight regulatory aspects of metabolic network behavior. Plant Physiol 142: 1380–1396 Caspar T, Huber SC, Somerville C (1985) Alterations in growth, photosynthesis, and respiration in a starch-less mutant of Arabidopsis thaliana (L.) deficient in chloroplast phosphoglucomutase activity. Plant Physiol 79: 11–17 Chia T, Thorneycroft SM, Chapple A, Messerli G, Chen J, Zeeman SC, Smith SM, Smith AM (2004) A cytosolic glucosyltransferase is required for the conversion of starch to sucrose in Arabidopsis leaves at night. Plant J 37: 853–863[CrossRef][Web of Science][Medline] Chritchley JH, Zeeman SC, Tahaka T, Smith AM, Smith SM (2001) A critical role for disproportionating enzyme in starch breakdown is revealed by a knock-out mutant in Arabidopsis. Plant J 26: 89–100[CrossRef][Web of Science][Medline] Davis BJ (1964) Disc electrophoresis. II. Method and application to human serum proteins. Ann N Y Acad Sci 121: 404–427[Web of Science][Medline] Deschamps P, Colleoni C, Nakamura Y, Suzuki E, Putaux JL, Buléon A, Haebel S, Ritte G, Steup M, Falcón LJ, et al (2008) Metabolic symbiosis and the birth of the plant kingdom. Mol Biol Evol 25: 536–548 Deuschle K, Chaudhuri B, Okumoto S, Lager I, Lalonde S, Frommer WB (2006) Rapid metabolism of glucose detected with FRET glucose nanosensors in epidermal cells and intact roots of Arabidopsis RNA-silencing mutants. Plant Cell 18: 2314–2325 Eckermann N, Fettke J, Steup M (2002) Identification of polysaccharide binding proteins by affinity electrophoresis in inhomogeneous polyacrylamide gels and subsequent SDS-PAGE/matrix-assisted laser desorption ionization-time of flight analysis. Anal Biochem 304: 180–192[CrossRef][Web of Science][Medline] Edner C, Li J, Albrecht T, Mahlow S, Hejazi M, Hussein H, Kaplan FI, Guy C, Smith SM, Steup M, et al (2007) Glucan, water dikinase activity stimulates breakdown of starch granules by plastidial beta-amylases. Plant Physiol 145: 17–28 Farre EM, Tiessen A, Roessner U, Geigenberger P, Trethewey RN, Willmitzer L (2001) Analysis of the compartmentation of glycolytic intermediates, nucleotides, sugars, organic acids, amino acids, and sugar alcohols in potato tubers using a nonaqueous fractionation method. Plant Physiol 127: 685–700 Fernie AR, Roessner U, Geigenberger P (2001) The sucrose analog palatinose leads to a stimulation of sucrose degradation and starch synthesis when supplied to discs of growing potato tubers (Solanum tuberosum). Plant Physiol 125: 1967–1977 Fernie AR, Swiedrych A, Tauberger E, Lytovchenko A, Trethewey RN, Willmitzer L (2002a) Potato plants exhibiting combined antisense repression of cytosolic and plastidial isoforms of phosphoglucomutase surprisingly approximate wild type with respect to the rate of starch synthesis. Plant Physiol Biochem 40: 921–927[CrossRef][Web of Science] Fernie AR, Tauberger E, Lytovchenko A, Roessner U, Willmitzer L, Trethewey RN (2002b) Antisense repression of cytosolic phosphoglucomutase in potato (Solanum tuberosum L.) results in severe growth retardation, reduction in tuber number and altered carbon metabolism. Planta 214: 510–520[CrossRef][Web of Science][Medline] Fettke J, Chia T, Eckermann N, Smith AM, Steup M (2006) A transglucosidase necessary for starch degradation and maltose metabolism in leaves acts on cytosolic heteroglycans (SHG). Plant J 46: 668–684[CrossRef][Web of Science][Medline] Fettke J, Eckermann N, Kötting O, Ritte G, Steup M (2007) Novel starch-related enzymes and carbohydrates. Cell Mol Biol 52: OL883–OL904[Medline] Fettke J, Eckermann N, Poeste S, Pauly M, Steup M (2004) The glycan substrate of the cytosolic (Pho 2) phosphorylase isoform from Pisum sativum L.: identification, linkage analysis and subcellular localization. Plant J 39: 933–946[CrossRef][Web of Science][Medline] Fettke J, Eckermann N, Tiessen A, Geigenberger P, Steup M (2005a) Identification, subcellular localization and biochemical characterization of water-soluble heteroglycans (SHG) in the leaves of Arabidopsis thaliana L.: distinct SHG reside in the cytosol and in the apoplast. Plant J 43: 568–585[CrossRef][Web of Science][Medline] Fettke J, Poeste S, Eckermann N, Tiessen A, Pauly M, Geigenberger P, Steup M (2005b) Analysis of cytosolic heteroglycans from leaves of transgenic potato (Solanum tuberosum L.) plants that under- or overexpress the Pho 2 phosphorylase isozyme. Plant Cell Physiol 46: 1987–2004 Fulton DC, Stettler M, Mettler T, Vaughan CK, Li J, Francisco P, Gil M, Reinhold H, Eicke S, Messerli G, et al (2008) β-Amylase4, a noncatalytic protein required for starch breakdown, acts upstream of three active β-amylases in Arabidopsis chloroplasts. Plant Cell 20: 1040–1058 Geigenberger P, Stitt M, Fernie AR (2004) Metabolic control analysis and regulation of the conversions of sucrose to starch in growing potato tubers. Plant Cell Environ 27: 655–673[CrossRef] Geiger M, Stitt M, Geigenberger P (1998) Metabolism in slices from growing potato tubers responds differently to addition of sucrose and glucose. Planta 206: 234–244[CrossRef][Web of Science] Giege P, Heazlewood JL, Roessner-Tunali U, Millar AH, Fernie AR, Leaver CJ, Sweetlove LJ (2003) Enzymes of glycolysis are functionally associated with the mitochondrion in Arabidopsis cells. Plant Cell 15: 2140–2151 Graham JW, Williams TC, Morgan M, Fernie AR, Ratcliffe RG, Sweetlove LJ (2007) Glycolytic enzymes associate dynamically with mitochondria in response to respiratory demand and support substrate channeling. Plant Cell 19: 3723–3738 Hejazi M, Fettke J, Haebel S, Edner C, Paris O, Frohberg C, Steup M, Ritte G (2008) Glucan, water dikinases phosphorylates crystalline maltodextrins and thereby initiates solubilization. Plant J 55: 323–334[CrossRef][Web of Science][Medline] Holtgrawe D, Scholz A, Altmann B, Scheibe R (2005) Cytoskeleton-associated, carbohydrate-metabolizing enzymes in maize identified by yeast two-hybrid screening. Physiol Plant 125: 141–156[CrossRef] Kofler H, Häusler RE, Schulz B, Gröner F, Flügge UI, Weber A (2000) Molecular characterisation of a new mutant allele of the plastid phosphoglucomutase in Arabidopsis, and complementation of the mutant with the wild-type cDNA. Mol Gen Genet 263: 978–986[CrossRef][Web of Science][Medline] Kosegarten H, Mengel K (1994) Evidence for a glucose 1-phosphate translocator in storage tissue amyloplasts of potato (Solanum tuberosum) suspension-cultured cells. Physiol Plant 91: 111–120[CrossRef] Lu Y, Sharkey TD (2004) The role of amylomaltase in maltose metabolism in the cytosol of photosynthetic cells. Planta 207: 271–274 Lytovchenko A, Bieberich K, Willmitzer L, Fernie AR (2002a) Carbon assimilation and metabolism in potato leaves deficient in plastidial phosphoglucomutase. Planta 215: 802–811[CrossRef][Web of Science][Medline] Lytovchenko A, Hajirezaei M, Eickmeier I, Mittendorf V, Sonnewald U, Willmitzer L, Fernie AR (2005a) Expression of an Escherichia coli phosphoglucomutase in potato (Solanum tuberosum L.) results in minor changes in tuber metabolism and a considerable delay in tuber sprouting. Planta 221: 915–927[Medline] Lytovchenko A, Schauer N, Willmitzer L, Fernie AR (2005b) Tuber-specific cytosolic expression of a bacterial phosphoglucomutase in potato (Solanum tuberosum L.) dramatically alters carbon partitioning. Plant Cell Physiol 46: 588–597 Lytovchenko A, Sweetlove LJ, Pauly M, Fernie AR (2002b) The influence of cytosolic phosphoglucomutase on photosynthetic carbohydrate metabolism. Planta 215: 1013–1021[CrossRef][Web of Science][Medline] Naeem M, Tetlow IJ, Emes MJ (1997) Starch synthesis in amyloplasts purified from developing potato tubers. Plant J 11: 1095–1103[CrossRef][Web of Science] Niittylä T, Messerli G, Trevisan M, Chen J, Smith AM, Zeeman SC (2004) A previously unknown maltose transporter essential for starch degradation in leaves. Science 303: 87–89 Nothnagel EA (1997) Proteoglycans and related components in plant cells. Int Rev Cytol 174: 195–291[Web of Science][Medline] Reiter WD (2008) Biochemical genetics of nucleotide sugar interconversion reactions. Curr Opin Plant Biol 11: 236–243[CrossRef][Web of Science][Medline] Ritte G, Lloyd JR, Eckermann N, Rottmann A, Kossmann J, Steup M (2002) The starch-related R1 protein is an Scheible WR, Pauly M (2004) Glycosyltransferases and cell wall biosynthesis: novel players and insights. Curr Opin Plant Biol 7: 285–295[CrossRef][Web of Science][Medline] Scheidig A, Fröhlich A, Schulze S, Lloyd JR, Kossmann J (2002) Down regulation of a chloroplast-targeted β-amylase leads to a starch-excess phenotype in leaves. Plant J 30: 581–591[CrossRef][Web of Science][Medline] Smith AM, Stitt M (2007) Coordination of carbon supply and plant growth. Plant Cell Environ 30: 1126–1149[CrossRef][Medline] Smith AM, Zeeman SC, Smith SM (2005) Starch degradation. Annu Rev Plant Biol 56: 73–97[CrossRef][Medline] Steichen JM, Petty RV, Sharkey TD (2008) Domain characterization of a 4- Steup M (1990) Starch degrading enzymes. In PJ Lea, ed, Methods in Plant Biochemistry, Vol 3. Academic Press, London, pp 103–128 Steup M, Latzko E (1979) Intracellular localization of phosphorylases in spinach and pea leaves. Planta 145: 69–75[CrossRef][Web of Science] Takamiya S, Fukui T (1978) Purification and multiple forms of phosphoglucomutase from potato tubers. Plant Cell Physiol 19: 759–768 Tauberger E, Fernie AR, Emmermann M, Renz A, Kossmann J, Willmitzer L, Trethewey RN (2000) Antisense inhibition of plastidial phosphoglucomutase provides compelling evidence that potato tuber amyloplasts import carbon from the cytosol in the form of glucose-6-phosphate. Plant J 23: 43–53[Medline] Viola R, Roberts AG, Haupt S, Gazzani S, Hancock RD, Marmiroli N, Machray GC, Oparka KJ (2001) Tuberization in potato involves a switch from apoplastic to symplastic phloem unloading. Plant Cell 13: 385–398 Weber A, Servaites JC, Geiger DR, Kofler H, Hille D, Gröner F, Hebbeker U, Flügge I (2000) Identification, purification, and molecular cloning of a putative plastidic glucose transporter. Plant Cell 12: 787–801 Weise SE, Weber APM, Sharkey TD (2004) Maltose is the major form of carbon exported from chloroplasts at night. Planta 218: 474–482[CrossRef][Web of Science][Medline] Yariv J, Rapport M, Graf L (1962) The interaction of glycosides and saccharides with antibody to the corresponding phenylazo glycoside. Biochem J 85: 383–388[Web of Science][Medline] Zeeman SC, Delatte T, Messerli G, Umhang M, Stettler M, Mettler T, Streb S, Reinhold H, Kötting O (2007a) Starch breakdown: recent discoveries suggest distinct pathways and novel mechanisms. Funct Plant Biol 34: 465–473[CrossRef] Zeeman SC, Smith SM, Smith AM (2007b) The diurnal metabolism of leaf starch. Biochem J 401: 13–28[CrossRef][Web of Science][Medline] Zhang L, Häusler RE, Greiten C, Hajirezaei MR, Haferkamp I, Neuhaus HE, Flügge UI, Ludewig F (2008) Overriding the co-limiting import of carbon and energy into tuber amyloplasts increases the starch content and yield of transgenic potato plants. Plant Biotechnol J 6: 453–464[CrossRef][Medline] This article has been cited by other articles:
|
|||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| ASPB Publications | PLANT PHYSIOLOGY® | THE PLANT CELL | |
|---|---|---|---|
TOP
ABSTRACT
RESULTS
