JXB Advance Access originally published online on March 26, 2004
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Journal of Experimental Botany, Vol. 55, No. 400, pp. 1231-1244, May 1, 2004
© 2004 Oxford University Press
Photosynthetic Carbon Fluxes |
Using mutants to probe the in vivo function of plastid envelope membrane metabolite transporters
Received 12 September 2003; Accepted 18 December 2003
Michigan State University, Department of Plant Biology, East Lansing, MI, 48824, USA
* To whom correspondence should be addressed. Fax: +1 517 432 5294. E-mail: aweber{at}msu.edu
 |
Abstract |
|---|
 Top Abstract IntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
During the last 15 years, much progress has been made in discovering genes encoding solute transporters of the inner plastid envelope membrane. For example, genes encoding transporters for phosphorylated intermediates, dicarboxylates, adenine nucleotides, inorganic anions, and monosaccharides have been cloned. In many cases, the corresponding proteins have been expressed in recombinant host systems for further functional studies, thus allowing detailed in vitro characterization of transporter properties. Knowledge of the gene sequences encoding these transporters have allowed reverse-genetic approaches to study transporter function in vivo. Antisense repression and T-DNA insertion mutagenesis have provided a range of transgenic and mutant plants in which the activity of specific plastid envelope transporters are massively decreased or abolished. Plants with altered transporter activities represent excellent tools to probe the in vivo function of these transporters. Moreover, changing the permeability of the plastid envelope membrane permits the targeted manipulation of subcellular metabolite pools.
Key words: Antisense, envelope membrane, gene knockout, membrane transport, metabolite transport, mutant, plastid, translocator.
|
Introduction |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
Plastids are the metabolic factories of plant cells. A range of biosynthetic pathways, such as the light and carbon reactions of photosynthesis, ammonia assimilation, starch biosynthesis, sulphate assimilation, fatty acid biosynthesis, tocopherol biosynthesis, the biosynthesis of terpenoids, and the shikimic acid pathway are exclusively or predominantly localized in plastids. However, plastid metabolism is intertwined with metabolism in the cytosol and precursors, intermediates, and end-products of plastid-localized biosynthetic pathways need to be exchanged with the surrounding cytosol. This tangled metabolic network requires massive traffic of solutes across the plastid envelope membrane (Fig. 1). Two bilayer membranes, the inner and the outer plastid envelope membrane separate the plastid stroma from the cytosol. It is generally accepted that the inner plastid envelope membrane represents the actual permeability barrier between plastid and cytosol, whereas the outer envelope membrane is freely permeable for solutes up to a molecular weight of approximately 10 kDa (Flügge, 2000; Flügge and Benz, 1984; Heldt and Sauer, 1971; Heldt et al., 1972). However, this view has recently been challenged by the discovery of specific, regulated solute pores in the outer envelope membrane (Bölter and Soll, 2001; Soll et al., 2000).
|
The study of metabolite transport across the plastid envelope membrane only became possible after reliable methods for the isolation of intact chloroplasts had been established in the early 1960s (Walker, 1964). Initially, the effect of externally applied solutes on photosynthetic carbon dioxide assimilation and oxygen evolution was studied. Later, the efflux of solutes from chloroplasts was measured (Bassham et al., 1968). The direct measurement of the uptake of radiolabelled substrates into intact, isolated chloroplasts by the silicon-oil layer filtration centrifugation technique opened the way to a detailed characterization of the kinetic properties of a range of metabolite transporters in the chloroplast inner envelope membrane (Heldt, 2002; Heldt and Rapley, 1970). These biochemical studies were instrumental for understanding the metabolite flux between plastid and cytosol. However, it was not until 1989 that the first cDNA sequence of a plastid envelope membrane transporter became available (Flügge et al., 1989). The establishment of recombinant expression systems for these transporters and of methods for reconstituting recombinant metabolite transporters into artificial lipid vesicles allowed detailed in vitro studies of transporter properties (Loddenkötter et al., 1993). Although the recently finished sequence of the Arabidopsis genome indicates the existence of some 150 putative metabolite transporters in the plastid envelope membrane (Ferro et al., 2002; Koo and Ohlrogge, 2002; Schwacke et al., 2003), up to now only a small number of these transporters have been identified at the molecular level (Table 1). For the majority of these proteins, their functions have been confirmed by in vitro studies using reconstituted, recombinant proteins, or by complementation of yeast knockout mutants.
|
Although these studies provided a wealth of information on the biochemistry and molecular biology of plastid envelope membrane solute transporters, they were unable to resolve the in vivo role of these proteins. This only became possible after new methods emerged that allowed the modulation or the elimination of transporter activity in intact plants. For example, the antisense repression and the constitutive overexpression of TPT in transgenic tobacco plants enabled a comprehensive analysis of the control coefficient that TPT exerts on photosynthetic carbon dioxide assimilation (Häusler et al., 2000a, b).
Several recent reviews have addressed the physiology, identification, and functional characterization of metabolite transporters in the inner envelope plastid membrane (Fischer and Weber, 2002; Flügge, 1998, 1999; Flügge et al., 2003; Neuhaus and Wagner, 2000; Weber and Flügge, 2002). This review focuses on recent results from knockdown and knockout studies of plastid envelope membrane solute transporters.
|
Common features of all inner plastid envelope membrane transporters |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
All characterized solute transporters of the inner plastid envelope membrane are nuclear-encoded polytopic membrane proteins. The precursor proteins are synthesized on cytosolic ribosomes and are post-translationally imported into the inner plastid envelope membrane. Although the exact pathway for membrane insertion and processing has not been solved yet, it is believed that the precursor proteins are processed to the mature forms by a stromal processing peptidase after or during insertion into the inner envelope membrane.
|
The plastidic phosphate translocator family |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
The plastidic phosphate translocator family consists of four members, namely TPT, PPT, GPT, and XPT. The cDNAs encoding these proteins have been expressed in yeast cells and the recombinant proteins have been reconstituted into liposomes so that their substrate specificities and kinetic parameters could be determined in vitro (Flügge, 1999; Flügge et al., 2003). The plastidic phosphate translocator family is a subgroup of a larger, distantly related protein family of drug and sugar transporters of mostly unknown function that is found in plants, animals, and fungi (Fischer and Weber, 2002; Knappe et al., 2003a). Until now, no related proteins have been detected in prokaryotes, raising the interesting question of the evolutionary origin of this protein family. In contrast to the originally proposed model that functional mature phosphate translocators consist of dimers of two subunits with six transmembrane helices segments each, newer results indicate that the subunits of the dimer possess seven to eight
-helical membrane-spanning segments (Knappe et al., 2003a). Proteins of the individual subfamilies share 35–50% amino acids identity, whereas the members of the same subfamily from different plant species share 75–90% identical amino acids (Flügge et al., 2003; Knappe et al., 2003a).
The triose phosphate/phosphate translocator (TPT)
The TPT exports the end-products of photosynthetic carbon dioxide assimilation, the triose phosphates glyceraldehyde-3-phosphate (GAP) and dihydroxyacetone-3-phosphate (DAP), from the chloroplast stroma to the cytosol (Fig. 2). Under physiological conditions, this transport occurs in strict counter-exchange with ortho-phosphate (Pi) or 3-phosphoglyceric acid (3-PGA). TPT transcripts are confined to photosynthetically active tissues in tobacco and Arabidopsis. In wild-type plants, triose phosphate export from the plastid to the cytosol by the TPT represents the principal pathway for the export of recently assimilated carbon dioxide and the predominant source of carbon skeletons for sucrose biosynthesis (Flügge, 1999).
|
Antisense repression of TPT in potato and tobacco: Given this important role in carbon allocation and partitioning, a reduction in TPT activity in intact plants was to be expected to result in a dramatic phenotype. This hypothesis was tested by antisense repression of TPT expression in transgenic potato plants (Riesmeier et al., 1993). Surprisingly, a reduction of the TPT transcript to almost undetectable levels and a corresponding reduction in TPT protein and activity by more than 30% compared with the wild type did not lead to an apparent phenotype. However, a detailed analysis of starch and sucrose levels in these transgenic plants, throughout the photoperiod, revealed that the lack in TPT transport capacity was compensated by an increased allocation of recently assimilated carbon dioxide into the plastidic starch pool and a corresponding decrease in soluble sugars during the light period. During the following dark period, starch breakdown and sucrose export from leaves was increased compared with wild-type plants. Hence, the reduced TPT activity resulted in a change in carbon partitioning between the plastidic starch and cytosolic sucrose pools during the light period and increased export of reduced carbon from source to sink tissues during the night (Heineke et al., 1994).
Antisense-TPT tobacco plants follow a different strategy. In these plants, triose phosphate translocator activity was reduced by approximately 70% compared with the wild type (Häusler et al., 1998). Using pulse–chase studies and analysis of starch and soluble sugars throughout the photoperiod, it was demonstrated that tobacco plants, displaying reduced capacity for triose phosphate export from plastids, allocate more of the recently fixed carbon dioxide into the starch pool. By contrast with antisense-TPT potato plants, in tobacco, the increased starch biosynthesis rate during the light period was accompanied by increased starch breakdown in the light, increased activity of starch degrading enzymes, an increase in glucose export capacity from chloroplasts, and by increased hexokinase activity (Häusler et al., 1998). Thus, recently assimilated carbon, instead of being exported to the cytosol in the form of triose phosphates, is shuttled through the transitory starch pool to form glucose (and most likely also maltose) that is exported from the plastid stroma to the cytosol by specific transporters, thereby bypassing the bottleneck generated by the reduction in TPT activity. A detailed control analysis demonstrated that TPT activity did not limit photosynthetic carbon dioxide assimilation under ambient conditions. However, maximal rates of carbon dioxide assimilation were severely reduced in antisense-TPT tobacco plants at saturating CO2 and light conditions and increased in plants showing overexpression of TPT. This indicates that wild-type levels of TPT activity can limit CO2 assimilation under saturating conditions (Häusler et al., 2000a, b).
A T-DNA insertion mutant for TPT: Recently Arabidopsis T-DNA insertion mutants in the TPT gene (tpt-1) became available (Schneider et al., 2002). Although TPT is a single-copy gene in the Arabidopsis genome, tpt-1 did not display an apparent phenotype under standard growth conditions. Similar to the transgenic tobacco plants described above, the lack in TPT activity in tpt-1 was compensated by an increased flux of recently assimilated carbon through the starch pool in the light. Inhibition of starch biosynthesis in tpt-1, by crossing it to a mutant in the large subunit of the plastidic ADP-glucose pyrophosphorylase (AGPase), led to a dramatic dwarfish phenotype, thereby demonstrating that starch biosynthesis is the major salvage pathway for the compensation of a TPT knockout. Surprisingly, a genetic cross of tpt-1 to the starch excess mutant sex1 caused only a minor growth inhibition and a moderate phenotype. Sex1 is deficient in the Arabidopsis homologue of the potato protein R1 (Yu et al., 2001), a starch–water dikinase that transfers phosphate groups from ATP to starch (Ritte et al., 2002). It was demonstrated that R1 activity is required for the turnover of transitory starch in Arabidopsis and of transitory and storage starch in potato (Lorberth et al., 1998; Yu et al., 2001). Interestingly, the abundance of a water-soluble polydextran was increased in the tpt-1/sex1 double-mutant, indicating the presence of a highly mobile maltodextrin pool that may serve as a temporary carbon store in the absence of starch turnover (Schneider et al., 2002).
These examples nicely underline the plasticity of plant metabolism and, in addition, illustrate the co-ordination of plastidic starch biosynthesis with cytosolic sucrose biosynthesis by phosphate and 3-PGA levels in the plastid stroma. The results obtained with the transgenic tobacco plants and the Arabidopsis T-DNA insertion mutants clearly demonstrate that transitory starch can be simultaneously synthesized and degraded in the light. Moreover, these mutants and transgenic plants indicate that the export of triose phosphates from chloroplasts does not play a role during night-time export of reduced carbon resulting from the degradation of transitory starch. This is in agreement with the finding that a loss-of-function mutant in the cytosolic fructose 1,6-bisphosphatase was unable to synthesize sucrose from triose phosphates in the light period, but sucrose synthesis was detectable in the dark (Micallef et al., 1996a, b; Micallef and Sharkey, 1996).
Furthermore, the Arabidopsis TPT knockout mutant represents a valuable tool to study critical components of starch biosynthesis and turnover, because the knockout of non-redundant components in starch metabolism will lead to a dramatic phenotype in the TPT knockout background, as demonstrated by the AGPase/TPT double knockout.
The phosphoenolpyruvate/phosphate translocator (PPT)
Recombinant, reconstituted PPT catalyses the strict counter-exchange of phosphoenolpyruvate (PEP) with Pi or 2-PGA in vitro. Under physiological conditions, it is believed to operate as a PEP/Pi-antiporter (Fischer et al., 1997). Chloroplasts, unlike most non-green plastids, lack the activities of enolase and/or phosphoglyceromutase (Bagge and Larsson, 1986; Borchert et al., 1993; Van der Straeten et al., 1991); hence triose phosphates cannot be converted to PEP in the plastid stroma. Therefore PPT is required for the import of PEP into the plastid stroma of C3-plants where it serves as the precursor for the shikimate pathway and, in oil-storing seeds, potentially also as a source of both carbon and ATP for fatty acid biosynthesis (Fig. 3). In cauliflower and maize, PPT transcripts were detected in green and non-green tissue, with steady-state transcript levels being highest in non-green tissues (Fischer et al., 1997). The Arabidopsis genome encodes two PPT-proteins, AtPPT1 and AtPPT2 (Knappe et al., 2003a). These genes are differentially expressed throughout the plant: AtPPT1 is expressed in the vasculature of leaves and roots, with the highest expression in xylem parenchyma cells, but not in leaf mesophyll cells, whereas AtPPT2 is expressed ubiquitously in leaves, but not in roots (Knappe et al., 2003b).
|
Chloroplasts of C4-type plants contain a very active PPT (Gross et al., 1990; Huber and Edwards, 1977; Rumpho et al., 1988). By contrast with C3-type plants, where PEP is imported into plastids by PPT, the main function of PPT in C4 mesophyll chloroplasts is the export of PEP from chloroplasts to the cytosol where it serves as a precursor for the formation of OAA from PEP and bicarbonate in a reaction catalysed by PEPC. Although a PPT from maize has been reported (Fischer et al., 1997), its expression pattern (high in non-green tissues, low in leaves) argues against a function for this specific gene product in C4 photosynthesis.
PPT knockout mutants: A knockout mutant in PPT1, cue1, displays a reticulate phenotype in which interveinal regions of the leaves are visibly pale, whereas paraveinal regions are green. This phenotype is accompanied by aberrantly shaped mesophyll cells containing abnormal chloroplasts, whereas bundle sheath cells and chloroplasts appear like the wild type (Streatfield et al., 1999). The phenotype was attributed to a reduced import of PEP into plastids, thus reducing of the supply of precursors to the shikimate pathway, and possibly to other metabolic pathways as well, such as isopentenyl biosynthesis. This idea was supported by the observation that the contents of and the flux into metabolites derived from the shikimate pathway, such as phenylpropanoids, were severely reduced in cue1 (Streatfield et al., 1999; Voll et al., 2003). Additional support for the hypothesis that the cue1 phenotype is caused by a lack of PEP in the plastid stroma came from the observation that constitutive overexpression of pyruvate:phosphate dikinase (PPDK) in the stroma of cue1 was able to abrogate the reticulate phenotype of mutant (Voll et al., 2003). PPDK can generate PEP from pyruvate that can still be taken up from the cytosol, thereby bypassing the defective PPT1. This elegant approach also demonstrated that Arabidopsis mesophyll chloroplasts have a substantial capacity for pyruvate transport across the envelope membrane. However, the overexpression of PPDK did not alleviate all aspects of the cue1 phenotype, indicating that the phenotype cannot exclusively be attributed to a reduction in PEP transport capacity (Voll et al., 2003).
A recent study surprisingly showed that PPT1 expression is confined to the paraveinal leaf regions in the wild type (Knappe et al., 2003b). Thus, it is the region that appears normal in cue1 that specifically lacks the AtPPT1 gene product! A second PPT-gene (PPT2) is ubiquitously expressed in leaves, providing a basic constitutive transport capacity for PEP in the absence of PPT1 expression, as is the case in cue1. In contrast to AtPPT1, AtPPT2 expression is not detectable in roots. In the light of these results, is has been proposed that the vasculature-located AtPPT1 is involved in the generation of phenylpropanoid metabolism-derived signal molecules that trigger the development in interveinal leaf regions, and that this signal originates from the root vasculature where only AtPPT1 (but not AtPPT2) is expressed (Knappe et al., 2003b).
The glucose 6-phosphate/phosphate translocator (GPT)
The GPT has the broadest substrate specificity among the members of the plastidic phosphate translocator family (Flügge et al., 2003). The purified, reconstituted translocator protein is able to exchange radiolabelled Pi for triose phosphates, glucose 6-phosphate (Glc 6-P), the pentose phosphates ribulose 5-phosphate (Ru-5-P), ribose-5-phosphate (Rib 5-P), and xylulose 5-phosphate (Xul 5-P), as well as PEP, 3-PGA, and erythrose 4-phosphate. However, considering the kinetic constants for these substrates, GPT in vivo most likely operates as a Glc 6-P/Pi antiporter (Eicks et al., 2002; Flügge et al., 2003; Kammerer et al., 1998). The proposed function of GPT is the supply of non-green plastids with Glc 6-P as a precursor for starch biosynthesis and the oxidative pentose pathway (Fig. 4). Glc 6-P is the sole precursor for starch biosynthesis in Arabidopsis, which was demonstrated by the fact that a mutation in the plastidic isozyme of phosphoglucomutase (conversion of Glu 6-P to Glu 1-P) leads to a starch-free phenotype. Hence, in Arabidopsis, there exists no pathway for the uptake of Glc 1-P into plastids (Caspar et al., 1985; Kofler et al., 2000). A mutant in the plastidic isozyme of phosphoglucoisomerase (conversion of Fru 6-P to Glu 6-P) contains substantial amounts of starch in non-green tissues, whereas photosynthetically active tissues were found to be essentially starch-free (Yu et al., 2000). This finding indicates that mesophyll chloroplasts do not have significant Glc 6-P transport capacity that could bypass the interrupted link between the Calvin cycle and starch biosynthesis.
|
The Arabidopsis genome encodes two closely related GPT proteins (AtGPT1 and AtGPT2). No knockout mutants in either gene have been described in the literature, and preliminary evidence obtained in Flügge’s laboratory indicates that a knockout in AtGPT1 may be embryo lethal (A Schneider, personal communication).
The xylulose 5-phosphate/phosphate translocator XPT
The most recent addition to the plastidic PT family is the xylulose 5-phosphate/phosphate translocator XPT. This protein was identified by mining the Arabidopsis genome sequence for genes encoding proteins with similarities to other members of the PT family. It shares approximately 50% identical amino acids with AtGPT1 and AtGPT2 and about 35–40% amino acid sequence identity with other members of the PT family. A single copy gene encodes XPT in Arabidopsis (Eicks et al., 2002). XPT steady-state transcript levels were found to be similar in all tested tissues such as flowers, leaves, roots, and shoots. This is in contrast to the TPT that is exclusively expressed in photosynthetically active tissues. A detailed analysis using a promoter–GUS fusion demonstrated expression of the XPT gene in vegetative parts such as leaves of all developmental stages, roots, and particularly strong in root tips. GUS activity was restricted to sepals, filaments, the upper part of the style, and the stigma in floral tissue, but no activity was observed in petals, young ovaries, and the anthers. Strong GUS activity was obtained in the pods and seeds of developed fruits, but no staining was evident during the early fruit development period (Eicks et al., 2002).
Expression of the corresponding cDNA in yeast and reconstitution of the purified, recombinant protein into liposomes demonstrated that XPT accepts triose phosphates, Xul 5-P, Ru 5-P, and Ery 4-P as counter-exchange substrates for inorganic phosphate. Rib 5-P and hexose phosphates are not transported. The low Ki values for triose phosphates (0.4 mM) and Xul 5-P (0.8 mM) indicate that these compounds efficiently compete with Pi for binding to XPT. Hence, under physiological conditions, XPT most likely transports triose phosphates, Xul 5-P, and inorganic phosphate, whereas the transport of PEP, 3-PGA, Ery 4-P, and Ru 5-P is unlikely, taking into account the rather high concentrations necessary to achieve this counter-exchange (Eicks et al., 2002).
The proposed function of XPT is the anaplerotic supply of plastids with Xul 5-P under conditions where demand for Calvin cycle intermediates by metabolic pathways such as the shikimate pathway is high and withdrawal of Calvin cycle intermediates such as Ery 4-P would deplete carbon skeletons required for sustained regeneration of Ru 1,5-bP (Eicks et al., 2002). However, a closer look at the stoichiometry of the Calvin cycle throws some doubt on this proposal. Assuming a constant rate of carbon dioxide assimilation and Ery 4-P production by transketolase, a withdrawal of Ery 4-P for the entry reaction of the shikimate pathway would disturb the stoichiometry by which Ery 4-P can be used for the generation of sedoheptulose 1,7-bisphosphate by transaldolase. This disequilibrium would result in a relative increase in the Xul 5-P production rather than a demand for Xul 5-P. Since transketolase is confined to the stroma in Arabidopsis (Eicks et al., 2002), Ery 4-P cannot be produced in the cytosol and, consequently, a role of XPT in the import of Ery 4-P into chloroplasts can be ruled out.
An alternative hypothesis is proposed, that XPT is involved in supplying the plastidic oxidative pentose phosphate pathway (OPPP) with carbon precursors. This hypothesis is based on two observations: (i) Glc 6-P cannot be taken up into chloroplasts because chloroplasts do not possess a GPT (see above), and (ii) phosphorolytic degradation of transitory starch (as an alternative source for the generation of hexose-phosphates in the plastid stroma) does not occur at significant rates (Smith et al., 2003). Hence, an alternative pathway for feeding the plastidic OPPP would be required (Fig. 5): One molecule of Xul 5-P would be imported into the plastid stroma in counter-exchange with one molecule of Pi. Together with five molecules of pentose phosphates, Xul 5-P would enter the non-oxidative branch of the OPPP, giving rise to four molecules of Fru 6-P and two molecules of triose phosphate. The two molecules of triose phosphates would be converted to one molecule of Fru 6-P and one molecule of Pi by the combined action of aldolase and FbPase. The resulting five molecules of Fru 6-P are converted into Glc 6-P by phosphoglucose isomerase, giving rise to five molecules of Glc 6-P. These would enter the oxidative branch of the OPPP, yielding five molecules of CO2, five molecules of NADPH, and five molecules of pentose-phosphates. The Pi generated by the FBPase reaction can be exchanged for an additional Xul 5-P, leading into another round of the cycle. The low affinity of XPT for Rib 5-P and Ru 5-P avoids the counter-productive exchange of pentose phosphates with each other, thus providing an efficient pathway for Xul 5-P import.
|
Although this model provides an attractive hypothesis for the role of XPT in plastid types that lack GPT activity, it may not adequately reflect the situation in non-green tissues because GPT provides a direct pathway for Glc 6-P import in these tissues. As proposed previously (Eicks et al., 2002) an additional role of XPT may be the transport of peroxisome-derived pentose phosphates (resulting from peroxisomal G6PDH activity that is required for the peroxisomal ascorbate–glutathione cycle) into plastids.
Neither knockout mutants nor antisense plants for XPT have been reported to date. Preliminary evidence obtained in Flügge’s group indicates that an XPT T-DNA insertion mutant does not display an obvious phenotype (M Eicks, personal communication). This may be due to the partially redundant function of GPT that also accepts pentose phosphates as substrates, in which case double knockouts may be required to gain further insight into the physiological role of this transporter. In addition, XPT function may become limiting only under specific metabolic conditions, therefore further studies will be required to analyse the effects of a XPT knockout under a broad range of growth conditions.
|
Plastidic dicarboxylate translocators (DiTs) |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
In leaves of most C3-type angiosperm plant species, ammonia is predominantly assimilated by the plastid-localized GS/GOGAT cycle (Hirel and Lea, 2001). This pathway assimilates ammonia resulting from nitrate/nitrite reduction; however, in leaves, about 90% of its capacity is taken up by reassimilation of ammonia that is released from the photorespiratory carbon cycle (Hirel and Lea, 2001). The essential role of the GS/GOGAT-system for the reassimilation of ammonia resulting from the photorespiratory pathway was established by the conditional lethal phenotype (viable at high CO2, conditions that suppress photorespiration) of mutants defective in Fd-GOGAT (Kendall et al., 1986; Somerville and Ogren, 1980) or GS2 (Blackwell et al., 1987).
2-Oxoglutarate (2-OG), the precursor for primary ammonia assimilation is synthesized in the cytosol and/or in the mitochondria (Hodges, 2002; Lancien et al., 2000). In addition, 2-OG that is generated in peroxisomes by the photorespiratory pathway needs to be shuttled back to plastids. Hence, plastids depend on transporters (dicarboxylate transporters) that import 2-oxoglutarate into plastids and that export the product of ammonia assimilation, glutamate, to the cytosol (Fig. 6A). This transport process is achieved by a pair of malate-coupled metabolite transporters that work in concert: the 2-oxoglutarate/malate translocator (DiT1) imports 2-oxoglutarate into the plastid in counter-exchange with the export of malate; the glutamate/malate translocator (DiT2) exports glutamate to the cytosol, in counter-exchange with the import of malate. Hence, the import of 2-oxoglutarate and export of glutamate occurs without net malate transport (Fig. 6B). This two-translocator model was developed based on kinetic studies of 2-oxoglutarate, malate, and glutamate uptake into isolated plastids (Flügge et al., 1988; Woo et al., 1987; Yu and Woo, 1992a, b). Both, DiT1 and DiT2 have recently been identified at the molecular level (Renné et al., 2003; Taniguchi et al., 2002; Weber et al., 1995).
|
Plastidic dicarboxylate transporters are hydrophobic polytopic membrane proteins that most likely operate as monomers containing 12 transmembrane
-helical segments. In this respect, they differ from mitochondrial dicarboxylate transporters that are composed of two identical subunits with six transmembrane helices each (Laloi, 1999; Walker and Runswick, 1993). Although functionally very similar to mitochondrial dicarboxylate transporters, plastidic dicarboxylate transporters are not related to mitochondrial transporters at the DNA or amino acid sequence level, but they are closely related to a family of eubacterial di- and tricarboxylate transporters (Weber and Flügge, 2002). The phylogenetic relationships of plastidic dicarboxylate translocators and the two- and three-translocators models for the transport of dicarboxylic acids and glutamate in plastids have recently been reviewed (Weber and Flügge, 2002). Therefore the following discussion concentrates on recent findings obtained with knockout mutants in DiT1 and DiT2.
The 2-oxoglutarate/malate translocator DiT1
The main function of DiT1 in C3-type plants is the import of 2-oxoglutarate from the cytosol into the plastid stroma where it serves as the carbon backbone for ammonia assimilation by the GS/GOGAT system. The specificity of DiT1 for 2-oxoglutarate was confirmed by the functional reconstitution of yeast-expressed, recombinant DiT1 from spinach and Arabidopsis (Taniguchi et al., 2002; Weber et al., 1995). DiT1 accepts the dicarboxylates 2-OG, fumarate, succinate, and malate, but not the amino acids glutamate, aspartate, or glutamine as substrates (Taniguchi et al., 2002; Weber et al., 1995).
Surprisingly, a recent study found that recombinant, reconstituted DiT1 from Arabidopsis also has the function of an oxaloacetate/malate transporter (Taniguchi et al., 2002). Similar results were obtained with the corresponding protein from spinach (Renné et al., 2003). Oxaloacetate uptake into chloroplasts was first reported by Heldt (Heldt and Rapley, 1970) and a specific oxaloacetate/malate translocator was previously characterized in isolated, intact chloroplasts from spinach and maize (Hatch et al., 1984). The function of the oxaloacetate/malate shuttle in C3-plants is the export of reducing equivalents from the plastid to the cytosol (Heineke et al., 1991). In C4-plants such as maize, the oxaloacetate flux into mesophyll cell plastids occurs at the rate of CO2 assimilation: oxalocetate (OAA) is formed by carboxylation of PEP by PEPC in the cytosol of mesophyll cells, OAA is then transported into mesophyll cell chloroplasts, reduced to malate by NADP+-MDH, and malate is subsequently exported to the cytosol of mesophyll cells to be delivered to bundle sheath cell chloroplasts where it is oxidatively decarboxylated by malic enzyme to pyruvate and CO2. The oxaloacetate/malate transporter has not been previously identified at the DNA or protein levels; therefore, the finding that DiT1 is able to transport OAA in vitro is particularly interesting. However, the kinetic constants of the recombinant, reconstituted translocator protein, in particular, the strong inhibition of oxaloacetate transport by malate, do not accurately concord with those obtained previously for the high affinity oxaloacetate/malate transporter of isolated spinach and maize chloroplasts (Hatch et al., 1984; Renné et al., 2003). Further studies will be required to clarify a possible function of DiT1 as the plastidic oxaloacetate/malate transporter.
Knockout mutants and antisense plants for DiT1: Two independent T-DNA insertion mutants for DiT1 have been isolated in Taniguchi’s laboratory and the absence of DiT1 transcripts in both mutants has been demonstrated (Taniguchi et al., 2002). In this initial study the knockout did not reveal any apparent phenotype. However, an additional T-DNA insertion allele has been isolated in the authors’ laboratory, that, when grown at ambient conditions, shows significantly reduced growth compared with the wild type (J Schneidereit, APM Weber, unpublished results). Further studies of all mutant alleles will be required before a definite phenotype of a DiT1 gene knockout can be established.
Transgenic tobacco plants have also been generated showing antisense repression of DiT1 (J Schneidereit et al., unpublished results). These plants are characterized by bleached regions along the vasculature. In intermediate phenotypes, the interveinal regions remain green whereas in strong phenotypes, leaves are yellowish and partially deformed, plants show stunted growth caused by massively reduced internode length, and flowering is delayed by several weeks. The transgenic plants accumulate intermediates of the TCA cycle, 2-OG, ammonia, and glutamine, whereas the levels of glutamate and glutamate-derived amino acids such as proline are dramatically reduced. By contrast with the wild type, feeding of 2-OG to detached leaves did not stimulate the production of glutamate or decrease the glutamine contents in the transgenic plants, indicating a block between 2-OG and the formation of glutamate from glutamine and 2-OG by Fd-GOGAT. This finding clearly supports a role of DiT1 in supplying plastids with 2-OG as precursor for ammonia assimilation by GS/GOGAT.
The glutamate/malate translocator DiT2
According to the two-translocator model, the function of DiT2 is the export of glutamate from plastids to the cytosol (Flügge et al., 1988; Woo et al., 1987). Genes encoding the DiT2 proteins from spinach, tobacco, Flaveria trinervia, sorghum, and Arabidopsis have recently been identified (Renné et al., 2003; Taniguchi et al., 2002; Weber and Flügge, 2002). The Arabidopsis genome encodes two closely related DiT2 genes, DiT2.1 and DiT2.2. DiT2 from spinach, Flaveria, and DiT2.1 from Arabidopsis have been expressed in yeast cells and the substrate specificity of the recombinant DiT2 proteins was tested. The function of DiT2 as a general dicarboxylate translocator with specificity for 2-OG, glutamate, aspartate, fumarate, and succinate, but not citrate or glutamine was demonstrated (Renné et al., 2003; Taniguchi et al., 2002).
Knockout mutants for DiT2: An EMS generated Arabidopsis mutant showing a deficiency in plastidic dicarboxylate transport (dct) was isolated in a screen for photorespiratory mutants in the early 1980s (Somerville and Ogren, 1983). This mutant requires elevated CO2 levels for growth and it lacks a protein with an apparent molecular mass of 45 kDa in its chloroplast envelope membrane (Somerville and Somerville, 1985). A barley mutant deficient in plastidic dicarboxylate transport and showing a similar photorespiratory phenotype was described in the following year (Wallsgrove et al., 1986). Based on the two-translocator model, either DiT1 or DiT2 may cause this phenotype because these two translocators work in concert. To identify the defective gene in dct, all three DiT-related genes were sequenced in dct and a single base-pair exchange was found in DiT2.1, leading to the replacement of a small, uncharged glycine residue by a negatively charged glutamate residue. Transformation of dct with the corresponding cDNA from spinach cured the photorespiratory phenotype, thus unequivocally demonstrating that dct is deficient in DiT2.1 gene function (Renné et al., 2003). Additional T-DNA insertion alleles for DiT2.1 have been isolated in Taniguchi’s group and these alleles also display a photorespiratory phenotype (Taniguchi et al., 2002). Hence, DiT2.1 is absolutely required for the operation of the photorespiratory carbon cycle. The second DiT2-related gene in the Arabidopsis genome, DiT2.2, is obviously not able to compensate for a deficiency in DiT2.1 function. Several T-DNA insertion mutants for DiT2.2 have been isolated. None of the mutant alleles displays an apparent phenotype (J Schneidereit, APM Weber, unpublished results). Taniguchi’s group assayed the function of recombinant, reconstituted DiT2.2, but no transport activity for di- and tricarboxylic acids and glutamate was detectable (Taniguchi et al., 2002). Hence, the function of DiT2.2 in Arabidopsis remains unclear.
|
The plastidic adenylate transporter (AATP) |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
The adenine nucleotide translocator was the first plastidic metabolite translocator to be studied (Heldt, 1969). Twenty-eight years later, the corresponding cDNA was identified in Neuhaus’ group (Kampfenkel et al., 1995; Neuhaus et al., 1997). In non-photosynthetic plastids (and chloroplasts during the dark) this protein is crucial for the provision of ATP from the cytosol that is required to drive biosynthetic reactions such as protein, RNA, starch, and fatty acid biosynthesis.
By contrast with mitochondrial adenine nucleotide translocators that consist of two identical subunits containing six transmembrane domains each (Laloi, 1999; Walker and Runswick, 1993), the mature plastidic AATP is most probably active as a monomer composed of 12 -helical transmembrane domains. The primary structure of plastidic AATPs is not related to that of mitochondrial adenine nucleotide transporters. However, several closely related proteins have been identified in a number of bacteria, all of them being obligate intracellular species, Together, the bacterial and the plastidic proteins constitute a distinct family of non-mitochondrial nucleotide transporters (Winkler and Neuhaus, 1999). The highly interesting phylogenetic relationships between these proteins have recently been addressed in two publications (Amiri et al., 2003; Linka et al., 2003).
Antisense repression and overexpression of AATP in potato: The Arabidopsis genome encodes two AATPs (Möhlmann et al., 1998). Knockout mutants in Arabidopsis have hitherto not been reported, but transgenic potato plants showing antisense repression of the endogenous AATP and constitutive overexpression of AtAATP1 have been analysed in detail (Geigenberger et al., 2001; Tjaden et al., 1998). The antisense lines showed a massive decrease in tuber size and tuber morphology was severely altered to structures resembling ginger root. Not only was the total amount of tuber starch reduced, but its composition was also affected. Starch from antisense plants exhibited a significantly reduced amylose to amylopectin ratio. By contrast, transgenic potato plants overexpressing AtAATP1 showed a marked increase in starch content and the amylose to amylopectin ratio was higher than in the wild type (Tjaden et al., 1998). Further analysis showed that tubers from antisense plants contained increased free sugar, UDP-Glc, and hexose phosphates levels, whereas PEP, isocitrate, adenine and uridine nucleotides, and inorganic pyrophosphate levels were slightly decreased. By contrast, adenine and uridine nucleotides, and inorganic pyrophosphate levels were elevated in tubers from sense plants, whereas soluble sugars remained unaltered. The ADP-Glc content was reduced by 50% in antisense tubers, whereas its content was increased up to 2-fold in sense tubers. These results suggested a close interaction between plastidial adenylate transport and starch biosynthesis, implying that ADP-Glc pyrophosphorylase is ATP-limited in vivo (Geigenberger et al., 2001).
The reduction of starch contents in AATP antisense potato plants is accompanied by decreased susceptibility to pathogens such as Erwinia carotovora, Phytophthora infestans, and Alternaria solani. This increased pathogen resistance could not be attributed to the elevated glucose contents of the transgenic plants and grafting experiments demonstrated the presence of a signal that is generated in AATP1 rootstocks and that primes wild-type scions for potentiated activation of cellular defence responses in leaves (Conrath et al., 2003).
The studies with these transgenic plants indicate that AATP represents an interesting target for the genetic engineering of starch-storing organs such as potato tubers and, similar to the PPT knockout mutant cue1 and the DiT2 knockout mutant dct, they demonstrate that the activity of transporters in the plastid envelope membrane can limit biosynthetic pathways in the plastid stroma that depend on precursor supply from the cytosol.
|
The chloroplast phosphate transporter PHT2;1 |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
The phosphate transporter PHT2;1 shares similarity with Na+/Pi transporters from mammals and fungi and with H+/Pi transporters from bacteria. This protein was originally described (Daram et al., 1999) as a low affinity phosphate transporter of the plasma membrane of Arabidopsis. Compared with related proteins from other kingdoms, PHT2;1 carries a long N-terminal extension of approximately 100 amino acids. It was hypothesized that this extension may have a regulatory function (Daram et al., 1999). A later study, however, demonstrated that a PHT2;1-GFP fusion was targeted to chloroplasts when transiently expressed in Arabidopsis plants and that the N-terminal extension of PHT2;1 has the characteristics of a plastid targeting signal (Versaw and Harrison, 2002). In yeast, the GFP-fusion protein was targeted to mitochondria. If, however, the first 71 amino acids of PHT2;1, representing the putative plastid targeting signal, were deleted and the truncated protein was fused to GFP, the chimeric protein was targeted in yeast cells to the plasma membrane and to structures associated with the ER. The truncated PHT2;1 protein was able to reconstitute the growth of a phosphate-uptake deficient yeast mutant, thereby demonstrating that PHT2;1 has the function of a phosphate transporter (Versaw and Harrison, 2002).
PHT2;1 transcripts could be detected in leaves but not in roots, and steady-state transcript levels increased upon transition from dark to light and upon resupply of phosphate to phosphate-depleted plants (Versaw and Harrison, 2002).
A PHT2;1 knockout mutant: A PHT2;1 null-mutant (pht2;1-1) has been isolated from a T-DNA insertion mutagenized Arabidopsis population (Versaw and Harrison, 2002). Pht2;1-1 displays reduced growth compared with the wild type and the leaf phosphate content is reduced by >20% at high-phosphate growth conditions. At limiting phosphate conditions, the total phosphate content of pht2;1-1 was similar to wild type plants; however, the phosphate content of roots was massively reduced, whereas the shoot phosphate levels were markedly increased. This finding indicates that pht2;1-1 affects phosphate allocation at the whole plant level. Moreover, by contrast with the wild type, a redistribution of phosphate from older leaves to younger leaves under conditions of Pi deficiency was not observed in pht2;1-1, underlining the role of PHT2;1 in the allocation and partitioning of phosphate at the whole plant level (Versaw and Harrison, 2002). It would be most interesting to study the partitioning of phosphate between plastids and cytoplasm in pht2;1-1 since it is very likely that a deficiency in PHT2;1 will lead to marked changes in subcellular phosphate pools in leaves.
The predominant expression of PHT2;1 in leaves argues against a significant role in starch biosynthesis in non-green tissues. Starch biosynthesis in most non-green tissues (with the exception of some cereal endosperm tissues (Denyer et al., 1996)) is fuelled by the import of Glc 6-P and ATP from the cytosol. ATP is stoichometrically exchanged for ADP, consequently, each imported and hydrolysed ATP also leads to the net import of one phosphate. Hence, phosphate would accumulate during starch biosynthesis if no mechanism exists for the unidirectional export of phosphate from plastids. Unidirectional transport of phosphate has been reported for isolated cauliflower bud plastids (Neuhaus and Maass, 1996), however, a gene encoding this transporter has not yet been identified. The Arabidopsis genome encodes several transporters with similarity to mammalian type I Na+/Pi phosphate transporters, some of which carry N-terminal extensions that have features of plastid targeting signals (Versaw and Harrison, 2002). It is tempting to speculate that one of these putative phosphate transporters may have the function of a plastidic phosphate exporter in non-photosynthetic cells.
|
Concluding remarks |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
Several additional genes encoding plastidic transporters, such as a putative glucose transporter (Weber et al., 2000), a sulphate transporter (Takahashi et al., 1999), a putative nitrite transporter (Rexach et al., 2000), an ADP-Glc transporter (Cao et al., 1995; Shannon et al., 1998; Sullivan and Kaneko, 1995), and others have been described in recent years. The identity of the vast majority of transporters, however, remains unknown. For example, transporters for amino acids, pyruvate, intermediates of the photorespiratory pathway such as glycolate and glycerate, metals, and many other substances are required. Systematic approaches, such as mining of plant genomes sequences for putative plastidic transporters (Koo and Ohlrogge, 2002; Schwacke et al., 2003), phenotypic analysis of corresponding knockout mutants, and functional analysis of recombinant candidate proteins will certainly lead to the discovery of novel transporters.
|
Acknowledgements |
|---|
We appreciate the communication of unpublished results by Anja Schneider and Michael Eicks, and of manuscripts prior to publication by Ulf-Ingo Flügge (University of Cologne, Germany). The helpful comments of the two anonymous reviewers are highly appreciated, in particular with respect to the in vivo role of XPT. A DAAD graduate fellowship to JS and a DFG postdoctoral fellowship to LMV is gratefully acknowledged. Work in the authors’ laboratory was funded by an MSU IRGP grant and by the National Science Foundation.
|
References |
|---|
|
TopAbstractIntroductionCommon features of all...The plastidic phosphate...Plastidic dicarboxylate...The plastidic adenylate...The chloroplast phosphate...Concluding remarksReferences |
|---|
Amiri H, Karlberg O, Andersson SE. 2003. Deep origin of plastid/parasite ATP/ADP translocases. Journal of Molecular Evolution 56, 137–150.[CrossRef][Web of Science][Medline]
Bagge P, Larsson C. 1986. Biosynthesis of aromatic amino acids by highly purified spinach chloroplasts—compartmentation and regulation of the reactions. Physiologia Plantarum 68, 641–647.
Bassham JA, Kirk M, Jensen RG. 1968. Photosynthesis by isolated chloroplasts. I. Diffusion of labeled photosynthetic intermediates between isolated chloroplasts and suspending medium. Biochimica et Biophysica Acta 153, 211–218.[Medline]
Blackwell RD, Murray AJS, Lea PJ. 1987. Inhibition of photosynthesis in barley with decreased levels of chloroplastic glutamine synthetase activity. Journal of Experimental Botany 38, 1799–1809.
Bölter B, Soll J. 2001. Ion channels in the outer membranes of chloroplasts and mitochondria: open doors or regulated gates? EMBO Journal 20, 935–940.[CrossRef][Web of Science][Medline]
Borchert S, Harborth J, Schünemann D, Hoferichter P, Heldt HW. 1993. Studies of the enzymic capacities and transport properties of pea root plastids. Plant Physiology 101, 303–312.[Abstract]
Cao HP, Sullivan TD, Boyer CD, Shannon JC. 1995. Bt1, a structural gene for the major 39–44 kDa amyloplast membrane polypeptides. Physiologia Plantarum 95, 176–186.[CrossRef]
Caspar T, Huber SC, Somerville C. 1985. Alterations in growth, photosynthesis, and respiration in a starchless mutant of Arabidopsis thaliana (L.) deficient in chloroplast phospho glucomutase activity. Plant Physiology 79, 11–17.
Conrath U, Linke C, Jeblick W, Geigenberger P, Quick WP, Neuhaus HE. 2003. Enhanced resistance to Phytophthora infestans and Alternaria solani in leaves and tubers, respectively, of potato plants with decreased activity of the plastidic ATP/ADP transporter. Planta 217, 75–83.[Web of Science][Medline]
Daram P, Brunner S, Rausch C, Steiner C, Amrhein N, Bucher M. 1999. Pht2;1 encodes a low-affinity phosphate transporter from Arabidopsis. The Plant Cell 11, 2153–2166.
Denyer K, Dunlap F, Thorbjornsen T, Keeling P, Smith AM. 1996. The major form of ADP-glucose pyrophosphorylase in maize endosperm is extra-plastidial. Plant Physiology 112, 779–785.[Abstract]
Eicks M, Maurino V, Knappe S, Flügge UI, Fischer K. 2002. The plastidic pentose phosphate translocator represents a link between the cytosolic and the plastidic pentose phosphate pathways in plants. Plant Physiology 128, 512–522.
Ferro M, Salvi D, Riviere-Rolland H, Vermat T, Seigneurin-Berny D, Grunwald D, Garin J, Joyard J, Rolland N. 2002. Integral membrane proteins of the chloroplast envelope: Identification and subcellular localization of new transporters. Proceedings of the National Academy of Sciences, USA 99, 11487–11492.
Fischer K, Kammerer N, Gutensohn M, Arbinger B, Weber A, Häusler RE, Flügge UI. 1997. A new class of plastidic phosphate translocators: a putative link between primary and secondary metabolism by the phosphoenolpyruvate/phosphate antiporter. The Plant Cell 9, 453–462.[Abstract]
Fischer K, Weber A. 2002. Transport of carbon in non-green plastids. Trends in Plant Science 7, 345–351.[CrossRef][Web of Science][Medline]
Flügge UI. 1998. Metabolite transporters in plastids. Current Opinion in Plant Biology 1, 201–206.[CrossRef][Web of Science][Medline]
Flügge UI. 1999. Phosphate translocators in plastids. Annual Review of Plant Physiology and Plant Molecular Biology 50, 27–45.[CrossRef][Web of Science][Medline]
Flügge UI. 2000. Transport in and out of plastids: does the outer envelope membrane control the flow? Trends in Plant Science 5, 135–377.[CrossRef][Web of Science][Medline]
Flügge UI, Benz R. 1984. Pore-forming activity in the outer membrane of the chloroplast envelope. FEBS Letters 169, 85–89.
Flügge UI, Fischer K, Gross A, Sebald W, Lottspeich F, Eckerskorn C. 1989. The triose phosphate-3-phosphoglycerate-phosphate translocator from spinach chloroplasts: nucleotide sequence of a full-length cDNA clone and import of the in vitro synthesized precursor protein into chloroplasts. EMBO Journal 8, 39–46.[Web of Science][Medline]
Flügge UI, Häusler RE, Ludewig F, Fischer K. 2003. Functional genomics of phosphate antiport systems of plastids. Physiologia Plantarum 118, 475–482.[CrossRef]
Flügge UI, Woo KC, Heldt HW. 1988. Characteristics of 2-oxoglutarate and glutamate transport in spinach chloroplasts. Studies with a double-silicone-layer centrifugation technique and in liposomes. Planta 174, 534–541.[CrossRef]
Geigenberger P, Stamme C, Tjaden J, Schulz A, Quick PW, Betsche T, Kersting HJ, Neuhaus HE. 2001. Tuber physiology and properties of starch from tubers of transgenic potato plants with altered plastidic adenylate transporter activity. Plant Physiology 125, 1667–1678.
Gross A, Brueckner G, Heldt HW, Flügge UI. 1990. Comparison of the kinetic properties, inhibition and labeling of the phosphate translocators from maize and spinach mesophyll chloroplasts. Planta 180, 262–271.
Hatch MD, Dröscher L, Flügge UI, Heldt HW. 1984. A specific translocator for oxaloacetate transport in chloroplasts. FEBS Letters 178, 15–19.
Häusler RE, Schlieben NH, Flügge UI. 2000a. Control of carbon partitioning and photosynthesis by the triose phosphate/phosphate translocator in transgenic tobacco plants (Nicotiana tabacum L.). II. Assessment of control coefficients of the triose phosphate/phosphate translocator. Planta 210, 383–390.[CrossRef][Web of Science][Medline]
Häusler RE, Schlieben NH, Nicolay P, Fischer K, Fischer KL, Flügge UI. 2000b. Control of carbon partitioning and photosynthesis by the triose phosphate/phosphate translocator in transgenic tobacco plants (Nicotiana tabacum L.). I. Comparative physiological analysis of tobacco plants with antisense repression and overexpression of the triose phosphate/phosphate translocator. Planta 210, 371–382.[CrossRef][Web of Science][Medline]
Häusler RE, Schlieben NH, Schulz B, Flügge UI. 1998. Compensation of decreased triose phosphate/phosphate translocator activity by accelerated starch turnover and glucose transport in transgenic tobacco. Planta 204, 366–376.[CrossRef][Web of Science][Medline]
Heineke D, Kruse A, Flügge UI, Frommer WB, Riesmeier JW, Willmitzer L, Heldt HW. 1994. Effect of antisense repression of the chloroplast triose-phosphate translocator on photosynthetic metabolism in transgenic potato plants. Planta 193, 174–180.
Heineke D, Riens B, Grosse H, Hoferichter P, Peter U, Flügge UI, Heldt HW. 1991. Redox transfer across the inner chloroplast envelope membrane. Plant Physiology 95, 1131–1137.
Heldt HW. 1969. Adenine nucleotide translocation in spinach chloroplasts. FEBS Letters 5, 11–14.[CrossRef][Web of Science][Medline]
Heldt HW. 2002. Three decades in transport business: studies of metabolite transport in chloroplasts—a personal perspective. Photosynthesis Research 73, 265–272.[CrossRef][Web of Science][Medline]
Heldt HW, Rapley L. 1970. Specific transport of inorganic phosphate, 3-phosphoglycerate and dihydroxyactone phosphate and of dicarboxylates across the inner membrane of spinach chloroplasts. FEBS Letters 10, 143–148.[CrossRef][Web of Science][Medline]
Heldt HW, Sauer F. 1971. The inner membrane of the chloroplast envelope as the side of specific metabolite transport. Biochimica et Biophysica Acta 234, 83–91.[Medline]
Heldt HW, Sauer F, Rapley L. 1972. Differentiation of the permeability properties of the two membranes of the chloroplast envelope. In: Forti G, Junk NV, eds. Proceedings of the International Congress of Photosynthesis Research, Vol. 2. The Hague, The Netherlands: 1345–1355.
Hirel B, Lea PJ. 2001. Ammonia assimilation. In: Lea PJ, Morot-Gaudry JF, eds. Plant nitrogen. Berlin: Springer-Verlag, 79–100.
Hodges M. 2002. Enzyme redundancy and the importance of 2-oxoglutarate in plant ammonium assimilation. Journal of Experimental Botany 53, 905–916.
Huber SC, Edwards GE. 1977. Transport in C4 mesophyll chloroplasts: evidence for an exchange of inorganic phosphate and phosphoenolpyruvate. Biochimica et Biophysica Acta 462, 603–612.[Medline]
Kammerer B, Fischer K, Hilpert B, Schubert S, Gutensohn M, Weber A, Flügge UI. 1998. Molecular characterization of a carbon transporter in plastids from heterotrophic tissues: the glucose6-phosphate/phosphate antiporter. The Plant Cell 10, 105–117.
Kampfenkel KH, Möhlmann T, Batz O, van Montague M, Inzé D, Neuhaus HE. 1995. Molecular characterization of an Arabidopsis thaliana cDNA encoding a novel putative adenylate translocator of higher plants. FEBS Letters 374, 351–355.[CrossRef][Web of Science][Medline]
Kendall AC, Wallsgrove RM, Hall NP, Turner JC, Lea PJ. 1986. Carbon and nitrogen-metabolism in barley (Hordeum vulgare L.) mutants lacking ferredoxin-dependent glutamate synthase. Planta 168, 316–323.[CrossRef]
Knappe S, Flügge UI, Fischer K. 2003a. Analysis of the plastidic phosphate translocator gene family in Arabidopsis and identification of new phosphate translocator-homologous transporters, classified by their putative substrate-binding site. Plant Physiology 131, 1178–1190.
Knappe S, Löttgert T, Schneider A, Voll L, Flügge UI, Fischer K. 2003b. Characterization of two functional phosphoenol pyruvate/phosphate translocator (PPT) genes in Arabidopsis: AtPPT1 may be involved in the provision of signals for correct mesophyll development. The Plant Journal 36, 411–420.[CrossRef][Web of Science][Medline]
Kofler H, Häusler RE, Schulz B, Gröner F, Flügge UI, Weber A. 2000. Molecular characterization of a new mutant allele of the plastid phosphoglucomutase in Arabidopsis, and complement ation of the mutant with the wild-type cDNA. Molecular and General Genetics 263, 978–986.
Koo AJ, Ohlrogge JB. 2002. The predicted candidates of Arabidopsis plastid inner envelope membrane proteins and their expression profiles. Plant Physiology 130, 823–836.
Laloi M. 1999. Plant mitochondrial carriers: an overview. Cell and Molecular Life Sciences 56, 918–944.
Lancien M, Gadal P, Hodges M. 2000. Enzyme redundancy and the importance of 2-oxoglutarate in higher plant ammonium assimilation. Plant Physiology 123, 817–824.
Linka N, Hurka H, Lang FB, Burger G, Winkler HH, Stamme C, Urbany C, Seil I, Kusch J, Neuhaus HE. 2003. Phylogenetic relationships of non-mitochondrial nucleotide transport proteins in bacteria and eukaryotes. Gene 306, 27–35.[CrossRef][Web of Science][Medline]
Loddenkötter B, Kammerer B, Fischer K, Flügge UI. 1993. Expression of the functional mature chloroplast triose phosphate translocator in yeast internal membranes and purification of the histidine-tagged protein by a single metal-affinity chromato graphy step. Proceedings of the National Academy of Sciences, USA 90, 2155–2159.
Lorberth R, Ritte G, Willmitzer L, Kossmann J. 1998. Inhibition of a starch-granule-bound protein leads to modified starch and repression of cold sweetening. Nature Biotechnology 16, 473–477.[CrossRef][Web of Science][Medline]
Micallef BJ, Dennis DT, Sharkey TD. 1996a. How do plants of Flaveria linearis having no activity of the ‘essential’ enzyme cytosolic fructose bisphosphatase survive? Plant Physiology 111, 298–298.
Micallef BJ, Sharkey TD. 1996. Genetic and physiological characterization of Flaveria linearis plants having a reduced activity of cytosolic fructose-1,6-bisphosphatase. Plant, Cell and Environment 19, 1–9.[Medline]
Micallef BJ, Vanderveer PJ, Sharkey TD. 1996b. Responses to elevated CO2 of Flaveria linearis plants having a reduced activity of cytosolic fructose-1,6-bisphosphatase. Plant, Cell and Environment 19, 10–16.[Medline]
Möhlmann T, Tjaden J, Schwoppe C, Winkler HH, Kampfenkel K, Neuhaus HE. 1998. Occurrence of two plastidic ATP/ADP transporters in Arabidopsis thaliana L.: molecular characterisation and comparative structural analysis of similar ATP/ADP translocators from plastids and Rickettsia prowazekii. European Journal of Biochemistry 252, 353–359.[Web of Science][Medline]
Neuhaus HE, Maass U. 1996. Unidirectional transport of orthophosphate across the envelope of isolated cauliflower-bud amyloplasts. Planta 198, 542–548.[Web of Science]
Neuhaus HE, Thom E, Möhlmann T, Steup M, Kampfenkel K. 1997. Characterization of a novel eukaryotic ATP/ADP translocator located in the plastid envelope of Arabidopsis thaliana L. The Plant Journal 11, 73–82.[CrossRef][Web of Science][Medline]
Neuhaus HE, Wagner R. 2000. Solute pores, ion channels, and metabolite transporters in the outer and inner envelope membranes of higher plant plastids. Biochimica et Biophysica Acta 1465, 307–323.[Medline]
Renné P, Dreßen U, Hebbeker U, Hille D, Flügge UI, Westhoff P, Weber APM. 2003. The Arabidopsis mutant dct is deficient in the plastidic glutamate/malate translocator DiT2. The Plant Journal 35, 316–331.[CrossRef][Web of Science][Medline]
Rexach J, Fernández E, Galván A. 2000. The Chlamydomonas reinhardtii Nar1 gene encodes a chloroplast membrane protein involved in nitrite transport. The Plant Cell 12, 1441–1453.
Riesmeier JW, Flügge UI, Schulz B, Heineke D, Heldt HW, Willmitzer L, Frommer WB. 1993. Antisense repression of the chloroplast triose phosphate translocator affects carbon partitioning in transgenic potato plants. Proceedings of the National Academy of Sciences, USA 90, 6160–6164.
Ritte G, Lloyd JR, Eckermann N, Rottmann A, Kossmann J, Steup M. 2002. The starch-related R1 protein is an alpha-glucan,water dikinase. Proceedings of the National Academy of Sciences, USA 99, 7166–7171.
Rumpho ME, Edwards GE, Yousif AE, Keegstra K. 1988. Specific labeling of the phosphate translocator in C3 and C4 mesophyll chloroplasts by tritiated dihydro-DIDS (1,2-ditritio-1,2-[2,2'-disulfo-4,4'-diisothiocyano] diphenylethane). Plant Physiology 86, 1193–1198.
Schneider A, Häusler RE, Kolukisaoglu U, Kunze R, van der Graaff E, Schwacke R, Catoni E, Desimone M, Flügge UI. 2002. An Arabidopsis thaliana knock-out mutant of the chloroplast triose phosphate/phosphate translocator is severely compromised only when starch synthesis, but not starch mobilisation is abolished. The Plant Journal 32, 685–699.[CrossRef][Web of Science][Medline]
Schwacke R, Schneider A, van der Graaff E, Fischer K, Catoni E, Desimone M, Frommer WB, Flügge UI, Kunze R. 2003. ARAMEMNON, a novel database for Arabidopsis integral membrane proteins. Plant Physiology 131, 16–26.
Shannon JC, Pien FM, Cao HP, Liu KC. 1998. Brittle-1, an adenylate translocator, facilitates transfer of extraplastidial synthesized ADP-glucose into amyloplasts of maize endosperms. Plant Physiology 117, 1235–1252.
Smith AM, Zeeman SC, Thorneycroft D, Smith SM. 2003. Starch mobilization in leaves. Journal of Experimental Botany 54, 577–583.
Soll J, Bölter B, Wagner R, Hinnah SC. 2000. .. response: the chloroplast outer envelope: a molecular sieve? Trends in Plant Science 5, 137–138.[CrossRef][Web of Science][Medline]
Somerville CR, Ogren WL. 1980. Inhibition of photosynthesis in Arabidopsis mutants lacking leaf glutamate synthase activity. Nature 286, 257–259.[CrossRef]
Somerville SC, Ogren WL. 1983. An Arabidopsis thaliana mutant defective in chloroplast dicarboxylate transport. Proceedings of the National Academy of Sciences, USA 80, 1290–1294.
Somerville SC, Somerville CR. 1985. A mutant of Arabidopsis deficient in chloroplast dicarboxylate transport is missing an envelope protein. Plant Science Letters 37, 217–220.[CrossRef]
Streatfield SJ, Weber A, Kinsman EA, Häusler RE, Li JM, Post-Beittenmiller D, Kaiser WM, Pyke KA, Flügge UI, Chory J. 1999. The phosphoenolpyruvate/phosphate translocator is required for phenolic metabolism, palisade cell development, and plastid- dependent nuclear gene expression. The Plant Cell 11, 1609–1621.
Sullivan TD, Kaneko Y. 1995. The maize brittle1 gene encodes amyloplast membrane polypeptides. Planta 196, 477–484.[Web of Science][Medline]
Takahashi H, Asanuma W, Saito K. 1999. Cloning of an Arabidopsis cDNA encoding a chloroplast localizing sulphate transporter isoform. Journal of Experimental Botany 50, 1713–1714.
Taniguchi M, Taniguchi Y, Kawasaki M, Takeda S, Kato T, Sato S, Tabata S, Miyake H, Sugiyama T. 2002. Identifying and characterizing plastidic 2-oxoglutarate/malate and dicarboxylate transporters in Arabidopsis thaliana. Plant Cell Physiology 43, 706–717.
Tjaden J, Möhlmann T, Kampfenkel K, Henrich G, Neuhaus HE. 1998. Altered plastidic ATP/ADP-transporter activity influences potato (Solanum tuberosum L.) tuber morphology, yield and composition of tuber starch. The Plant Journal 16, 531–540.[CrossRef][Web of Science]
Van der Straeten D, Rodrigues-Pousada RA, Goodman HM, Van Montagu M. 1991. Plant enolase: gene structure, expression, and evolution. The Plant Cell 3, 719–735.
Versaw WK, Harrison MJ. 2002. A chloroplast phosphate transporter, PHT2;1, influences allocation of phosphate within the plant and phosphate-starvation responses. The Plant Cell 14, 1751–1766.
Voll L, Häusler RE, Hecker R, Weber A, Weissenbock G, Fiene G, Waffenschmidt S, Flügge UI. 2003. The phenotype of the Arabidopsis cue1 mutant is not simply caused by a general restriction of the shikimate pathway. The Plant Journal 36, 301–317.[CrossRef][Web of Science][Medline]
Walker DA. 1964. Improved rates of carbon dioxide fixation by illuminated chloroplasts. Biochemical Journal 92, 22C–23C.[Medline]
Walker JE, Runswick MJ. 1993. The mitochondrial transport protein superfamily. Journal of Bioenergetics and Biomembranes 25, 435–446.[CrossRef][Web of Science][Medline]
Wallsgrove RM, Kendall AC, Hall NP, Turner JC, Lea PJ. 1986. Carbon and nitrogen-metabolism in a barley (Hordeum vulgare L.) mutant with impaired chloroplast dicarboxylate transport. Planta 168, 324–329.[CrossRef]
Weber A, Flügge UI. 2002. Interaction of cytosolic and plastidic nitrogen metabolism in plants. Journal of Experimental Botany 53, 865–874.
Weber A, Menzlaff E, Arbinger B, Gutensohn M, Eckerskorn C, Flügge UI. 1995. The 2-oxoglutarate/malate translocator of chloroplast envelope membranes: molecular cloning of a transporter containing a 12-helix motif and expression of the functional protein in yeast cells. Biochemistry 34, 2621–2627.[CrossRef][Medline]
Weber A, Servaites JC, Geiger DR, Kofler H, Hille D, Gröner F, Hebbeker U, Flügge UI. 2000. Identification, purification, and molecular cloning of a putative plastidic glucose translocator. The Plant Cell 12, 787–801.
Winkler HH, Neuhaus HE. 1999. Non-mitochondrial ATP transport. Trends in Biochemical Sciences 24, 64–68.[CrossRef][Web of Science][Medline]
Woo KC, Flügge UI, Heldt HW. 1987. A two-translocator model for the transport of 2-oxoglutarate and glutamate in chloroplasts during ammonia assimilation in the light. Plant Physiology 84, 624–632.
Yu JW, Woo KC. 1992a. Ammonia assimilation and metabolite transport in isolated chloroplasts. I. Kinetic measurement of 2-oxoglutarate and malate uptake via the 2-oxoglutarate translocator in oat and spinach chloroplasts. Australian Journal of Plant Physiology 19, 653–658.
Yu JW, Woo KC. 1992b. Ammonia assimilation and metabolite transport in isolated chloroplasts. II. Malate stimulates ammonia assimilation in chloroplasts isolated from leaves of dicotyledonous but not monocotyledonous species. Australian Journal of Plant Physiology 19, 659–669.
Yu TS, Kofler H, Häusler RE, et al. 2001. The Arabidopsis sex1 mutant is defective in the R1 protein, a general regulator of starch degradation in plants, and not in the chloroplast hexose transporter. The Plant Cell 13, 1907–1918.
Yu TS, Lue WL, Wang SM, Chen J. 2000. Mutation of Arabidopsis plastid phosphoglucose isomerase affects leaf starch synthesis and floral initiation. Plant Physiology 123, 319–326.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  C. Oesterhelt, S. Klocke, S. Holtgrefe, V. Linke, A. P. M. Weber, and R. Scheibe Redox Regulation of Chloroplast Enzymes in Galdieria sulphuraria in View of Eukaryotic Evolution Plant Cell Physiol., September 1, 2007; 48(9): 1359 - 1373. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Bouvier, N. Linka, J.-C. Isner, J. Mutterer, A. P.M. Weber, and B. Camara Arabidopsis SAMT1 Defines a Plastid Transporter Regulating Plastid Biogenesis and Plant Development PLANT CELL, November 1, 2006; 18(11): 3088 - 3105. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Barbier, C. Oesterhelt, M. D. Larson, R. G. Halgren, C. Wilkerson, R. M. Garavito, C. Benning, and A. P.M. Weber Comparative Genomics of Two Closely Related Unicellular Thermo-Acidophilic Red Algae, Galdieria sulphuraria and Cyanidioschyzon merolae, Reveals the Molecular Basis of the Metabolic Flexibility of Galdieria sulphuraria and Significant Differences in Carbohydrate Metabolism of Both Algae Plant Physiology, February 1, 2005; 137(2): 460 - 474. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||