Identification of a novel heteroglycan-interacting protein, HIP 1.3, from Arabidopsis thaliana

Journal of Plant Physiology 168 (2011) 1415–1425

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Identification of a novel heteroglycan-interacting protein, HIP 1.3, from Arabidopsis thaliana Joerg Fettke a,b , Adriano Nunes-Nesi c,1 , Alisdair R. Fernie c , Martin Steup b,∗ a b c

Mass Spectrometry of Biopolymers, Institute of Biochemistry and Biology, University of Potsdam, Karl-Liebknecht-Str. 24-25, Building 20, 14476 Potsdam-Golm, Germany Institute of Biochemistry and Biology, Plant Physiology, University of Potsdam, Karl-Liebknecht-Str. 24-25, Building 20, 14476 Potsdam-Golm, Germany Max-Planck Institute of Molecular Plant Physiology, Am Mühlenberg 1, 14469 Potsdam-Golm, Germany

a r t i c l e

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Article history: Received 25 August 2010 Received in revised form 23 September 2010 Accepted 24 September 2010 Keywords: Arabidopsis thaliana Carbohydrate binding proteins Cytosolic heteroglycans Maltose metabolism Starch metabolism

a b s t r a c t Plastidial degradation of transitory starch yields mainly maltose and glucose. Following the export into the cytosol, maltose acts as donor for a glucosyl transfer to cytosolic heteroglycans as mediated by a cytosolic transglucosidase (DPE2; EC 2.4.1.25) and the second glucosyl residue is liberated as glucose. The cytosolic phosphorylase (Pho2/PHS2; EC 2.4.1.1) also interacts with heteroglycans using the same intramolecular sites as DPE2. Thus, the two glucosyl transferases interconnect the cytosolic pools of glucose and glucose 1-phosphate. Due to the complex monosaccharide pattern, other heteroglycan-interacting proteins (HIPs) are expected to exist. Identification of those proteins was approached by using two types of affinity chromatography. Heteroglycans from leaves of Arabidopsis thaliana (Col-0) covalently bound to Sepharose served as ligands that were reacted with a complex mixture of buffer-soluble proteins from Arabidopsis leaves. Binding proteins were eluted by sodium chloride. For identification, SDS-PAGE, tryptic digestion and MALDI-TOF analyses were applied. A strongly interacting polypeptide (approximately 40 kDa; designated as HIP1.3) was observed as product of locus At1g09340. Arabidopsis mutants deficient in HIP1.3 were reduced in growth and contained heteroglycans displaying an altered monosaccharide pattern. Wild type plants express HIP1.3 most strongly in leaves. As revealed by immuno fluorescence, HIP1.3 is located in the cytosol of mesophyll cells but mostly associated with the cytosolic surface of the chloroplast envelope membranes. In an HIP1.3-deficient mutant the immunosignal was undetectable. Metabolic profiles from leaves of this mutant and wild type plants as well were determined by GC–MS. As compared to the wild type control, more than ten metabolites, such as ascorbic acid, fructose, fructose bisphosphate, glucose, glycine, were elevated in darkness but decreased in the light. Although the biochemical function of HIP1.3 has not yet been elucidated, it is likely to possess an important function in the central carbon metabolism of higher plants. © 2010 Elsevier GmbH. All rights reserved.

Introduction Biosynthesis of transitory starch is essentially photosynthesisdriven and, therefore, restricted to the light phase. In the subsequent dark period, starch is mobilized to sustain metabolism and growth of the entire plant (Zeeman et al., 2007, 2010). The

Abbreviations: DMSO, dimethylsulfoxide; FITC, fluorescein isothiocyanate; HIP1.3, heteroglycan interacting protein No. 1.3; PMSF, phenylmethylsulfonyl fluoride; HPAEC-PAD, high performance anion exchange chromatography coupled to pulsed amperomeric detection; SHG, water soluble heteroglycan; SHGS , water soluble heteroglycan with an apparent size below 10 kDa; SHGL , water soluble heteroglycan with an apparent size of more than 10 kDa. ∗ Corresponding author. E-mail address: [email protected] (M. Steup). 1 Present address: Departamento de Biologia Vegetal, Universidade Federal de Vic¸osa, Vic¸osa, Minas Gerais, Brazil. 0176-1617/$ – see front matter © 2010 Elsevier GmbH. All rights reserved. doi:10.1016/j.jplph.2010.09.008

plastidial starch degrading path converts the storage polysaccharide mainly into neutral sugars (maltose and, to a minor extent, glucose) that both are exported into the cytosol (Zeeman et al., 2010). The export of maltose is mediated by the recently identified maltose transporter (designated as MEX; Niittylä et al., 2004) but, until now, the glucose transporter has only functionally been characterized (Schleucher et al., 1998; Weber, 2004). In the cytosol, maltose acts as glucosyl donor for a disproportionating reaction that is catalyzed by a transglucosidase (DPE2; Chia et al., 2004; Lu and Sharkey, 2004). As opposed to Arabidopsis, contradicting data have been reported on the intracellular location of DPE2 in Solanum tuberosum L. (Lloyd et al., 2004; Fettke et al., 2005b) but, recently, the cytosolic location has been confirmed (Lütken et al., 2010). DPE2 transfers a maltose-derived hexosyl residue to non-reducing ends of heteroglycans and releases the other glucosyl residue of the disaccharide (the one carrying the reducing end) as free glucose (Fettke et al., 2009). Arabidopsis mutants that are deficient

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in either a functional MEX or DPE2 are severely compromised in growth and possess a starch excess phenotype. In addition, they contain extremely high maltose levels but, depending on the target gene, the intracellular location of the disaccharide differs: In the MEX mutants most of the maltose is retained inside the chloroplast. By contrast, DPE2-deficient mutants possess high maltose levels both in the cytosol and the plastid (Lu and Sharkey, 2004) and, therefore, MEX appears to mediate a bidirectional maltose transport. Furthermore, the cytosolic heteroglycans from DPE2-deficient lines are both functionally and structurally altered (Fettke et al., 2006). However, the phenotypical similarities between the MEXand DPE2-deficient mutants strongly suggest that the plastidial starch–maltose conversion, the maltose export into the cytosol and the subsequent DPE2-mediated maltose metabolism constitute an essential carbon flux that cannot be functionally substituted by other routes. It should be noted that the cytosolic heteroglycans, despite their essential function in the cellular starch metabolism, do not strictly depend on a functional starch biosynthesis. Arabidopsis mutants that are unable to accumulate normal levels of leaf starch (such as lines deficient in the plastidial phosphoglucomutase isozyme; Casper et al., 1985) possess essentially unaltered levels of the cytosolic heteroglycans. These data concur with the fast labelling of the heteroglycans during photosynthetic 14 CO2 fixation and suggest an efficient flux between the operating reductive pentose phosphate cycle and the cytosolic polyglycans (Fettke et al., 2005a). Cytosolic heteroglycans mainly consist of arabinose and galactose and contain various minor compounds, such as fucose, rhamnose, and glucose. The number of glycosidic linkages exceeds 20 (Fettke et al., 2004, 2005a,b). Due to the selectivity of carbohydrate-active enzymes, it is reasonable to assume that many heteroglycan-interacting proteins exist. However, until now only two proteins are known that both act on terminal glucosyl residues of the heteroglycans, i.e. DPE2 and PHS2. In an attempt to identify additional heteroglycan-interacting proteins, an affinity chromatography was performed using a heteroglycan preparation covalently bound to a Sepharose gel. A complex mixture of buffersoluble proteins extracted from Arabidopsis leaves was passed through the affinity gel and polypeptides were selected that are desorbed by relatively high concentrations of sodium chloride. Following the identification of the major interacting protein, a functional analysis was performed including the subcellular localization and the phenotypical characterization of an Arabidopsis mutant lacking the functional target protein. Materials and methods Plant materials Plants (Arabidopsis thaliana Col-0) were grown under controlled long-day conditions (15 h light [200 ␮mol m−2 s−1 ; 22 ◦ C] and 9 h dark [18 ◦ C]; relative humidity 50% throughout the light–dark cycle). Arabidopsis mutants putatively deficient in HIP1.3 were obtained from NASC (http://arabidopsis.info). Potato plants (Solanum tuberosum L. var. Desiré) were grown under controlled conditions as previously described (Fettke et al., 2005b). SHG isolation Water-soluble heteroglycans were isolated and fractionated as previously described (Fettke et al., 2005a,b). SHG quantification For quantification, heteroglycans were hydrolyzed in 2 M trifluoro acetic acid (3 h at 100 ◦ C). The total monosaccharide content of

the hydrolysate was determined according to Waffenschmidt and Jaenicke (1987) using glucose as standard. Identification of HIP1.3 Two types of affinity gels were generated. One affinity gel was synthesized by coupling heteroglycans to epoxy-activated Sepharose 6B (Amersham Biosciences, Uppsala, Sweden; Procedure A). For the other affinity gel Fractogel EMD Amino(M) from Merck (Darmstadt, Germany) and partially oxidized heteroglycans were used (Procedure B). For both A and B, the entire chromatographic procedure consists of five steps: preparation of the affinity gel (a), affinity chromatography (b), SDS-PAGE (c), tryptic digestion of the target protein (d), and MALDI-TOF analysis of the resulting peptide mixtures (e). Procedure A (a) Preparation of the SHGT -containing affinity gels. SHGT (10–22 mg) was dissolved in 2.5 ml water and the pH value of the resulting solution was adjusted to 13 by adding NaOH. Epoxyactivated Sepharose 6B was pre-treated following the instructions of the producer. Subsequently, the epoxy-activated Sepharose (2.5 g each; resuspended in 2.5 ml water) was added to the alkaline polyglycan solution and the mixture was incubated for 22 h at 40 ◦ C under continuous agitation. Following coupling, the Sepharose gel was washed with water and residual activated groups were blocked with 1 M ethanolamine (10 ml; 5 h; 40 ◦ C). Deactivation was terminated by dialysis. The Sepharose gel was then washed with 0.5 M NaCl, dissolved in 0.1 M acetate buffer pH 4.0 and, subsequently, in 0.5 M NaCl in 0.1 M Tris–HCl pH 8.0 (40 ml each). In total, this two-step washing procedure was performed three times. Finally, the polyglycan-containing Sepharose was washed several times in water and was then stored at 4 ◦ C in 0.1% [w/v] sodium azide, dissolved in water. The heteroglycan containing gel and the control gel is designated as affinity gel AT and ACon , respectively. (b) Affinity chromatography. Following the covalent coupling of the respective glycans, the Sepharose gel (2.5 g fresh weight each) was filled into a small column and was equilibrated with precooled grinding buffer (100 mM Hepes-NaOH pH 7.5, 1 mM EDTA, 2 mM DTE, and 0.5 mM PMSF). Subsequently, Arabidopsis leaves were harvested in the middle of the light period and buffer-soluble proteins were extracted using grinding buffer. The extracted proteins (3 mg each; dissolved in grinding buffer) were loaded to the gel and non-interacting proteins were removed by washing with 10 ml buffer I (100 mM NaCl, dissolved in 100 mM Hepes-NaOH pH 7.5). Subsequently, interacting proteins were eluted by 1 M NaCl, dissolved in 100 mM Hepes-NaOH pH 7.5. During the entire affinity chromatography, temperature was kept at 4 ◦ C. Finally, the Sepharose gel was washed with 3 ml 8 M urea, dissolved in water, to ensure complete removal of proteins. Eluted proteins were concentrated by membrane filtration (10 kDa cut off) and the medium was replaced by grinding buffer (see above). Glycerol was added (final concentration 10% [v/v]) and the concentrated native proteins were stored at −80 ◦ C. Alternatively, proteins were denatured and used for SDS-PAGE. (c) SDS-PAGE. Proteins were denatured for 5 min at 95 ◦ C and SDS-PAGE was performed as previously described (Eckermann et al., 2002). Subsequently, proteins were visualized using either silver staining (Eckermann et al., 2002) or colloidal Coomassie RotiR -Blue (Roth, Karlsruhe, Germany) following the instructions of the producer. (d) Tryptic digestion. Polyacrylamide gel pieces were destained for approximately 1 h in a mixture of 40% [v/v] acetonitrile and 60% [v/v] 50 mM NH4 HCO3 . Following drying, modified trypsin

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(30 ng ␮l−1 ; Roche, Mannheim, Germany), dissolved in 50 mM NH4 HCO3 , was added and the mixture was incubated over night at 37 ◦ C. Peptides were stepwise extracted by acetonitrile, 5%[v/v] formic acid and, subsequently, by acetonitrile and were concentrated by lyophilisation. (e) MALDI-TOF analysis. ␣-Cyano-4-hydroxy cinnamic acid (15 mg ml−1 , dissolved in 70% [v/v] acetonitrile), served as matrix. A Reflex II MALDI-TOF (Bruker-Daltonik, Bremen, Germany) was used in the reflector mode. For analytes having a size of up to 3 kDa mass accuracy was higher than ±0.1 Da. For all identified proteins, sequence coverage was at least 30%. Procedure B (a) Preparation of the affinity gels carrying SHG subfractions. Subfractions I and II were prepared from SHGL by either field flow fractionation or precipitation with the ␤-Yariv reagent (for details see Fettke et al., 2005a,b). Subsequently, each subfraction (0.4–3 mg) was partially oxidized by incubation with 2 mM sodium periodate (40 min at room temperature; darkness). Oxidation was terminated by adding glycerol and exhaustive dialysis (MWCO 1 kDa; Spectrum Laboratories Inc., Rancho Dominguez, CA, USA) and the volume of the solutions was reduced by lyophilisation. The Fractogel was washed three times with water following the instructions of the producer. Subsequently, 1 ml each of the gel suspension was mixed with 300 ␮l of the oxidized glycans (0.4–1 mg each), 135 ␮l concentrated acetic acid and 200 ␮l sodium cyanoborohydride (60 mg ml−1 , dissolved in DMSO). The mixture was incubated for 6.5 h at 37 ◦ C under continuous agitation. Subsequently, sodium cyano borohydride (as above; 200 ␮l each) was added and incubation was continued for 24 h under otherwise identical conditions. The gels were then washed five times with water and were blocked over night in 1 M ethanolamine (30 ml) at room temperature. Finally, the affinity gels were treated and stored as described in procedure A. The subfraction I and subfraction II containing gels are designated affinity gel BCyt and affinity gel BApo , respectively. In some experiments, SHGL isolated from leaves of potato (S. tuberosum L.) was used as ligand. Coupling to the gel was done exactly as decribed for subfractions I and II. In these experiments, a control gel was used that lacks SHGL but otherwise was treated identically. The two gels are designated as affinity gel BT and BCon , respectively. (b) Affinity chromatography. Except were noted, chromatography was performed as described above (procedure A). (c)–(e) These steps were performed as described before (procedure A). Native PAGE and enzyme activity staining For native PAGE and enzyme staining of enzyme activities see Fettke et al. (2005a). Metabolic profiling Metabolites were analysed by GC-MS using a protocol that had been optimized for Arabidopsis (Lisec et al., 2006). Peaks were identified using TAGfinder (Luedemann et al., 2008) and spectral libraries housed in the Golm Metabolome Database (Kopka et al., 2005). Relative metabolite contents were calculated according to Roessner et al. (2001). Protein blotting For the production of an anti-HIP1.3 antibody, the partial sequence CEEDAVDPKSRHKGK was selected as hapten. The peptide was coupled to KLH and a polyclonal antipeptide antibody

Fig. 1. Heteroglycan-interacting proteins from Arabidopsis leaves as revealed by affinity chromatography and SDS-PAGE. The affinity gel consists of epoxy-activated gel, coupled to SHGT (affinity gel AT , a) or a SHGT -free control gel (affinity gel ACon , b; procedure A, see section “Materials and methods”). Buffer-soluble proteins (4 mg each) extracted from leaves of Arabidopsis wild type plants (Col-0) were loaded to the two gels. Non-interacting proteins were removed by washing. Subsequently, the eluates obtained with 1 ml buffer I were collected (lanes W). Interacting proteins were eluted with 1 M NaCl, dissolved in 100 mM Hepes-NaOH pH 7.5 (lanes E1). Finally the two gels were eluted with 8 M urea, dissolved in water (lanes E2). Eluted proteins were separated by 12% T SDS-PAGE. M: molecular weight markers. The white arrow marks the 42 kDa protein, designated as HIP1.3.

was raised in rabbits (Sigma-Genosys, Suffolk, UK). This antibody was also used for immunocytochemistry. Immuno blotting was performed as previously described (Fettke et al., 2005a,b). Immunocytochemistry Leaves from Arabidopsis wild type plants and the HIP1.3deficient mutant were harvested in the light and were processed essentially as described by Schächtele and Steup (1986). Results Identification of a novel heteroglycan interacting protein, HIP1.3 Heteroglycans prepared from leaves of Arabidopsis thaliana were covalently bound to Sepharose and, subsequently, the affinity gel was loaded with a complex mixture of buffer soluble proteins extracted from Arabidopsis leaves. Proteins that interact with the immobilized glycans were separated from all other polypeptides as they were retained during the washing procedure. Selectivity of the protein–carbohydrate interaction was ensured by coupling different heteroglycan (sub)fractions to the Sepharose matrix. In the first series of experiments, the total SHG preparation (SHGT ) was immobilized (affinity gel AT ). In this case, the carbohydrate preparation contains all glycans fulfilling the following criteria: they are heat-stable (95 ◦ C), extractable in 20% [v/v] ethanol, water-soluble and the apparent size is at least 1 kDa. As a control, SHGT was omitted and epoxy-activated Sepharose was immediately reacted with ethanolamine (affinity gel ACon ). Both columns were loaded with a mixture of buffer-soluble proteins extracted from leaves of A. thaliana Col-0 (4 mg each) and were then washed (see section “Materials and methods”) to remove all not or weakly binding proteins. Eluates obtained by a final washing step (1 ml each) were collected and analysed by SDS-PAGE to ensure quantitative removal of non-interacting proteins (Fig. 1a lane W).

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Fig. 2. Selective interaction of buffer-soluble proteins with either cytosolic (affinity gel BCyt ) or apoplastic (affinity gel BApo ) heteroglycans as revealed by affinity chromatography and SDS-PAGE. Equal amounts (3 mg each) of proteins extracted from wild type Arabidopsis leaves were loaded on the two affinity gels. Both gels were four times washed with buffer I containing 0.1 NaCl (lanes W), then twice with 0.2 M NaCl, dissolved in Hepes-NaOH pH 7.5 (lanes E1), and, finally three times with 1 M NaCl, dissolved in Hepes-NaOH pH 7.5 (lanes E2). Eluted proteins were separated by 7.5% T SDS-PAGE. The white arrow marks the 42 kDa protein, designated as HIP1.3.

Subsequently, the sodium chloride concentration was raised to 1 M and approximately 20 distinct protein bands were released among which a protein with an apparent molecular weight of approximately 40 kDa was prominent (Fig. 1a lane E1). Most of these proteins were undetectable in the corresponding eluate from the control column (Fig. 1b) and, therefore, protein–matrix interactions appear to be minor. Treatment of the two gels with 8 M urea had little effect (Fig. 1 lanes E2) indicating that 1 M NaCl allows for an almost complete elution of heteroglycan-interacting proteins. Using tryptic digestion, MALDI-TOF analyses of the resulting peptides and data base searching, the 40 kDa protein was identified as the product of gene At1g09340. This gene encodes a protein containing 379 amino acid residues equivalent to a mass of 42.6 kDa and a predicted IEP of 8.41. Due to the strong interaction with immobilized heteroglycans (see below) the protein was designated as heteroglycan-interacting protein1.3 (HIP1.3). When coupling non-derivatized heteroglycans to the epoxyactivated Sepharose (Procedure A; see section “Materials and methods”), the carbohydrates are covalently bound to the matrix only via the reducing ends and, therefore, the structure of the ligands is largely unchanged. If, however, the glycans used for immobilization represent a complex mixture of compounds differing in size (and, possibly, in reactivity) the coupling procedure used may, to some extent, be selective. In this case, the pattern of the covalently bound heteroglycans is expected to deviate from that of the carbohydrates applied in the synthesis of the affinity gel and the actual carbohydrate targets of the interacting proteins remain unknown. Thus, the data shown in Fig. 1 clearly indicate that, in principle, heteroglycan-interacting proteins can be identified by an affinity chromatographic separation. It is, however, uncertain whether or not these proteins interact with cytosolic heteroglycans (subfraction I), with the apoplastic glycans (subfraction II) or with both. Therefore, in the second series of experiments, three major changes were made. First, Fractogel-Amino(M)-Sepharose replaced epoxy-activated Sepharose. Secondly, either subfraction I or II was applied as ligand. Thirdly, gels were synthesized following procedure B which includes a moderate periodate-mediated oxidation of the glycans to be coupled. By this pre-treatment, some of the vicinal hydroxyl groups are converted to the reactive carbonyl function. Unavoidably, oxidation affects the ring structure of the respective glycosyl residue but forms additional coupling sites most of which are located in internal glycosyl residues rather than close to the reducing end of the glycan. Due to the stochastic nature of the oxidation, larger glycans are likely to contain more carbonyl groups and to be more efficiently coupled as compared to oligoglycans.

Furthermore, both subfractions I and II consist of highly branched polysaccharides (Fettke et al., 2005a) and, therefore, the majority of the glycan chains are expected to be unchanged if relatively few sugar residues are oxidized. Equal amounts of buffer-soluble proteins were loaded to the affinity gel BCyt and BApo . Proteins that did weakly or not interact with the immobilized ligand were removed from the column by repetitive washings using a 0.1 M NaCl containing buffer. As revealed by SDS-PAGE, a few proteins were only gradually removed from both affinity gels (Fig. 2 lanes W) suggesting weak interactions with either the matrix or with each of the two immobilized glycans. Because of this delayed elution, washing was continued until the amount of the proteins released strongly decreased. Subsequently, the two columns were treated with stepwise increasing concentrations of sodium chloride but each step was repeated at least once (Fig. 2). Using 0.2 M NaCl as eluent, the protein patterns liberated from the two affinity gels differed largely. The pattern eluted from affinity gel BCyt consisted of more than 10 protein species. By contrast, the protein pattern released from affinity gel BApo was less complex and contained approximately five bands (Fig. 2). As revealed by SDS-PAGE, none of these bands comigrated with any protein eluted from the subfraction I-containing gel. These data indicate that the major proteins recovered selectively interact with either cytosolic (subfraction I) or apoplastic (subfraction II) heteroglycans implying an insignificant interaction with the matrix. Interestingly, elution of HIP1.3 required a NaCl concentration higher than 0.2 M but the same elution procedure did not, to any noticeable extent, release HIP1.3 from the subfraction II-containing gel. Likewise, treatment with urea was inefficient (Fig. 2 lanes E2). From these data three conclusions are reached: First, HIP1.3 selectively binds to the cytosolic heteroglycans and, secondly, interaction is strong as liberation requires high concentrations of sodium chloride. Finally, high salt conditions elute relatively large quantities of HIP1.3 suggesting an interaction with a major carbohydrate structure within subfraction I. In another series of experiments, the NaCl level that is required for the liberation of HIP1.3 was determined more precisely. The minimum eluting concentration was found to be approximately 0.25 M (data not shown). In another experiment, heteroglycans were isolated from leaves of S. tuberosum L. Subsequently, the SHGL fraction (that contains both subfractions I and II) was treated with periodate and then coupled to the Fractogel-Amino(M)-Sepharose. HIP1.3 from Arabidopsis leaves did bind to the affinity gel BT but not to the control gel (affinity gel BCon ). For complete elution, a rel-

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Fig. 3. Interspecies interaction between HIP1.3 from Arabidopsis and heteroglycans from Solanum tuberosum as revealed by affinity chromatography followed by SDSPAGE. Buffer-soluble proteins (4 mg each) from Arabidopsis wild type leaves that had been harvested at the beginning of the light period were loaded on affinity gel BT (+SHGT ) and BCon (-SHGT ). The two gels were washed with buffer containing 0.1 M NaCl (lanes W), with 1 M NaCl, dissolved Hepes-NaOH pH 7.5 (lanes E1), and, finally, with 8 M urea, dissolved in water (lanes E2). Aliquots of the crude extract loaded on the two gels were also subjected to the 7.5% T SDS-PAGE (lanes RE). The white arrow marks the 42 kDa protein, designated as HIP1.3.

Expression pattern and subcellular location of HIP1.3

Fig. 4. Organ-specific expression of HIP1.3. Extracts from various Arabidopsis wild type organs were subjected to SDS-PAGE (9% T separation gel), followed by immuno blotting using the anti-HIP1.3 antibody (a; 5 ␮g protein each) or to native PAGE and stained for DPE2 (b) or phosphorylase (c) activity. In (b) and (c), the separation gel (7.5% T) contained 0.25% [w/v] glycogen from oyster. For enzyme activity staining, the separation gels were incubated over night at 37 ◦ C. R: roots, yL: young leaves, mL: mature leaves; Sh: shoot; S: siliques, Fl: flowers, Se: seeds.

For several reasons, the organ-specific expression and the subcellular distribution of HIP1.3 were carefully determined. First, any potential biochemical function of HIP1.3 within the central carbon metabolism suggests that the protein is strongly expressed in photosynthesis-competent tissues. Secondly, it is reasonable to assume that the selective and strong interaction of HIP1.3 with cytosolic heteroglycans as observed in vitro is potentially relevant only if in vivo the protein has direct access to the target glycans. Accessibility would be easily given if HIP1.3 resides in the same compartment as subfraction I, i.e. in the cytosol. Finally, for the same gene product a chloroplastic location has recently been predicted (Hassidim et al., 2007; see section “Discussion”). Therefore, an empirical confirmation (or correction) of the predicted subcellular location of HIP1.3 is urgently needed. Organ-specific expression. Expression of HIP1.3 was studied at the protein level. Buffer-soluble proteins were extracted from rosette leaves (having a length of at least 0.8 cm), from leaves of flowering plants (leaf length at least 3 cm), roots, shoots, siliques and seeds. Equal amounts of buffer-soluble proteins were loaded on a SDS-containing polyacrylamide gel and, following electrophoresis, HIP1.3 was quantified by immuno blotting (Fig. 4a). As primary antibody, a polyclonal antipeptide antibody was used. HIP1.3 was strongly expressed in young and mature leaves, weakly in flowers and undetectable in roots or siliques. Using the same extracts, the activity of the cytosolic disproportionating enzyme, DPE2, and the pattern of the phosphorylase isoforms were determined. DPE2 activity was detected in all extracts but the highest activity was found in extracts from leaves and flowers (Fig. 4b). In a glycogen-containing separation gel, phosphorylase activity was recovered in three distinct bands: one band remains on top of the separation gel and represents one state of the cytosolic (PHS2) phosphorylase (Fig. 4c, I). PHS2 also exists in a more mobile state (Fig. 4c, II; see also Fettke et al., 2005a). As compared to both PHS2 states, the plastidial phosphorylase isozyme (PHS1) is more mobile (Fig. 4c, III). The highest phosphorylase activity

was observed in extracts from leaves and flowers. Thus, in leaves HIP1.3 and the two cytosolic glucosyl transferases are strongly expressed. Subcellular location. Several attempts were made to localize HIP1.3 in leaves. First, affinity chromatography (procedure A) was performed using extracts from wild type leaves and chloroplasts isolated from the same batch of leaves. Following isolation, buffer-soluble proteins were extracted. Buffer-soluble proteins from both leaves and chloroplasts were balanced to equal chlorophyll contents of the starting materials and were subjected to chromatography using affinity gel AT . Subsequently, eluate fractions were analysed by SDS-PAGE (Fig. 5a). Assuming a plastidial location of HIP1.3, the chromatographic separation of both samples is expected to result in essentially the same amount of HIP1.3. However, the leaf extract yielded significantly more target protein (Fig. 5a lanes E). Likewise, immuno blotting experiments performed with both protein preparations (that again were balanced for equal chlorophyll content) yielded a stronger immuno signal for the leaf extract compared to that of the isolated chloroplasts (Fettke, 2006). These data suggest that less than 50% of HIP1.3 is recovered in the chloroplast preparation. They are consistent with the non-aqueous fractionation of leaf tissue indicating that HIP1.3 does not clearly co-separate with plastidial proteins (nor with markers of any other compartment as well; data not shown). For an unequivocal subcellular localization of HIP1.3, immunocytochemistry was performed with leaf tissue using both wild type plants (Fig. 5b–d) and the HIP1.3-deficient mutant (Fig. 5e and f). The red chlorophyll fluorescence was used as a chloroplast marker (Fig. 5b). FITC-dependent fluorescence of the same wild type cell was paratially associated with but not restricted to chloroplasts and occurs also unevently distributed in the cytosol (Fig. 5c). This is most obvious when the two fluorescence signals were merged (Fig. 5d). As a control, leaf sections from the HIP1.3-deficient Arabidopsis mutant were analysed using exactly the same conditions of

ative high concentration of NaCl was required (Fig. 3). Thus, the glycan-HIP1.3 interaction is also observed in an interspecies approach.

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Fig. 5. Subcellular localization of HIP1.3. (a) Affinity chromatography (procedure A) followed by SDS-PAGE of extracts of isolated chloroplasts (WC and EC ) and of leaves (WL and EL ). Both extracts were balanced to be equivalent to equal chlorophyll contents. WC and WL : eluate fractions obtained by washing; EC and EL : fractions eluted with sodium chloride. The white arrow marks the position of HIP1.3. (b–f) Light microscopical analyses of cryocuts from Arabidopsis leaves of wild type plants (b–d) and of the HIP1.3-deficient mutant (e–f). (b) Chlorophyll fluorescence of wild type leaves; (c) FITC-dependent fluorescence of the same cryocut region; (d) merge of the chlorophyll and FITC fluorescence; (e) FITC-dependent fluorescence of the HIP1.3-deficient mutant; (f) merge of the FITC- and chlorophyll fluorescence of the same region of the cryocut. Bar: 10 ␮m.

specimen processing and microscopical evaluation (Fig. 5e and f). FITC-dependent fluorescent was low and similar to that observed in preimmune controls or specimen lacking the treatment with primary antibodies.

Functional in vivo analysis of HIP1.3 Based on the subcellular localization of HIP1.3, the protein has direct access to the cytosolic heteroglycans but any biochemical

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Fig. 6. Phenotypical characterization of the HIP1.3-deficient Arabidopsis mutant. (a) A HIP1.3-deficient mutant (left) and a wild type plant (right) grown for 8 weeks under defined conditions. (b) Affinity chromatography (procedure A) of buffer-soluble proteins extracted from leaves of the HIP1.3-deficient mutant (hip1.3) and from the wild type control (wt; 570 ␮g each). For details see Fig. 1. (c) Immuno blotting of crude extracts isolated from wild type and HIP1.3-deficient mutant with an antibody raised again HIP1.3. The white arrow marks the position of HIP1.3.

function exerted by HIP1.3 is unknown. For a functional characterization of the target protein, we choose three approaches: first we intended to determine the macroscopical phenotype of an Arabidopsis mutant lacking HIP1.3. Secondly, we characterized the heteroglycans isolated from an HIP1.3 deficient mutant. Finally, metabolic profiling was performed with leaves from this mutant. Data were compared with those obtained with wild type plants grown alongside with the mutant. Three SALK-lines from Arabidopsis that putatively contain an insertion in At1g09340 were tested. Line SALK 02111748 was selected as it was found to be homozygous even in the T5 generation (Fettke, 2006). In the following, this line is referred to as HIP1.3-deficient mutant. When grown under controlled conditions, growth of the mutant was strongly reduced (Fig. 6a). As opposed to the wild type control, buffer-soluble proteins extracted from leaves of the mutant did not contain, to any noticeable extent, the HIP1.3 as revealed by affinity chromatography (procedure A; Fig. 6b). Similarly, immuno blotting performed with proteins from the mutant (and the wild type control as well) did not result in any staining in the 40 kDa region (Fig. 6c). At the end of the light period, the fresh-weight based starch content of leaves from the HIP1.3-deficient mutant was essentially the same as that of the wild type control. Similarly, during darkness massive starch degradation occurred both in the mutant and the wild type (Table 1). At the end of both the light and the dark period, leaves from the mutant contained slightly elevated levels of heteroglycans (SHGT ) and of SHGL (Table 1). For a more detailed analysis, the various heteroglycan (sub)fractions were separated and subjected to acid hydrolysis. Subsequently, the resulting Table 1 Contents of heteroglycans and starch in leaves from the HIP1.3-deficient Arabidopsis mutant and the wild type control. Leaves were harvested during the middle of the light (Light) or the dark (Dark) period. The total heteroglycan contents (SHGT ) were calculated per gram fresh weight (FH). The high molecular weight fraction of the heteroglycans (SHGL ) is given as % SHGL . n = 4.

Wt Light Wt Dark hip1.3 Light hip1.3 Dark

SHGT [␮g g−1 FW]

% SHGL

± ± ± ±

79.3 77.3 81.3 79.50

217.6 225.2 253.5 261.43

8.9 7.5 13.5 9.31

± ± ± ±

Starch [mg glc g−1 FW] 1.1 3.4 2.3 3.41

4.2 0.8 4.4 0.51

± ± ± ±

0.8 0.2 0.3 0.05

monosaccharide patterns were resolved by HPAEC-PAD (Fig. 7). The relative fucose content is consistently reduced in the heteroglycans derived from the mutant. This effect is observed in the low-molecular SHG fraction (SHGS ), in subfraction I and in subfraction II as well. By contrast, in the hydrolysates of water-insoluble cell-wall materials the fucosyl content was essentially unchanged (Fig. 7c). However, the HIP1.3-deficient mutant exhibits more general significant phenotypical differences. This is indicated by metabolic profiling performed with leaves harvested during the light or dark period (Table 2). In the light period, the levels of arginine, ascorbate and dehydroascorbate, citrate, malate, succinate, galactinol, glutamate, tyrosine, myo-inositol, isomaltose, the polyamines spermidine and putrescene all decreased in the mutant. By contrast, with the exception of glycerate which is yet further decreased, in darkness the contents of ascorbate, fructose, glucose and fructose 1,6-bisphosphate, glycine, hydroxyproline, raffinose, succinate, threonate and tryptophan increase in the HIP1.3-deficient mutant. This opposing effect which is apparent even in the same metabolites is intriguing and perhaps implies a different function of HIP1.3 under different trophic conditions. The nature of changes of the metabolites is also interesting with a high number of alterations (such as those in myo-inositol, ascorbate, and threonate) being in metabolic pathways which are intimately linked to sugar phosphate utilization whilst others are closely involved in energy converting processes.

Discussion In this study, a classical biochemical approach has been used to identify heteroglycan-interacting proteins. Two types of affinity gels were synthesized differing in the matrix, in the mode of coupling and the ligands. Epoxy-activated Sepharose was reacted with the total heteroglycan preparation, SHGT and all ligand molecules were covalently bound via the reducing end. The second affinity gel used in this study was loaded with either the cytosolic heteroglycans (i.e. subfraction I) or the apoplastic glycans (subfraction II) both possessing additional intramolecular coupling sites. All interacting proteins were eluted from the affinity gel by using higher concentrations of sodium chloride rather than by the respective ligand in a soluble state. This mode was preferred as the isola-

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Fig. 7. Monosaccharide patterns of various glycan (sub)fractions isolated from leaves of the HIP1.3-deficient mutant and from a wild type control. Leaves were harvested at the end of the light or dark period. From each hydrolysate 5 ␮g carbohydrates were applied to the HPAEC column. All chromatograms were normalised to galactose. Abbreviations: Fuc: fucose; Ara: arabinose; Rha: rhamnose; Gal: galactose; Glc: glucose; Xyl: xylose; Man: mannose.

tion of large quantities of heteroglycans or heteroglycan-derived subfractions is very time-consuming. The approach chosen in this study is unbiased but, obviously, has some limitations: first, it is restricted to soluble (or solubilised yet functional) proteins. Secondly, if complex protein mixtures are to be analysed very low abundant compounds are often difficult to detect. Third, proteins that exhibit a strong interaction with the immobilized ligands are preferentially detected whereas those that bind weakly are difficult to distinguish from non-interacting compounds. Carbohydrate-active enzymes usually possess a modular organization and frequently contain, in addition to the catalytic region, one or several non-catalytic carbohydratebinding module(s) (CBMs; Boraston et al., 2004; Shoseyov et al., 2006). The biochemical functions of CBMs are diverse: firstly, they may alter the structure of the carbohydrate by binding to the target (Vaaje-Kolstad et al., 2005). Secondly, they may also support the respective enzyme activity by various targeting and proximity effects. As revealed for cell wall-related glycans, the CBM of a given carbohydrate-active enzyme not necessarily binds to the very same carbohydrate the catalytic domain acts on. Instead, the CBM may also interact with a glycan that, under in vivo conditions, is consistently and closely associated with the carbohydrate substrate (Hervé et al., 2010). For affinity chromatography, the occurrence of a CBM has important implications. If the protein–carbohydrate interaction

of a given carbohydrate-active enzyme is largely determined by the CBM(s) it is likely to be separated by affinity chromatography even if neither the actual reaction catalyzed nor the mode of catalysis (such as a random or ordered mechanism) is known. However, both the structural and the functional diversity of the CBMs identified is large and their apparent Km values vary by several orders of magnitude (Christiansen et al., 2009). Obviously, the affinity chromatography used favours the detection of enzymes containing high-affinity CBMs. For these reasons we did not attempt to identify all heteroglycaninteracting proteins but rather focused on a single polypeptide, HIP1.3. Several lines of independent evidence indicate that the in vivo function of HIP1.3 is related to the cytosolic heteroglycans and that it is involved in the central carbon metabolism: • The protein strongly and selectively interacts with cytosolic heteroglycans (Fig. 1). • The enzyme resides in the cytosol, thus in the very same compartment as the target glycans (Fig. 5b–d). • An HIP1.3-deficient Arabidopsis mutant that is impaired in growth possesses altered water-soluble heteroglycans (Fig. 6a) but insoluble cell wall constituents were unaltered. • Metabolic profiles from leaves of this mutant deviate from those of the wild type control. Furthermore, the metabolic phenotype

J. Fettke et al. / Journal of Plant Physiology 168 (2011) 1415–1425

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Table 2 Metabolic profiling of HIP1.3-deficient leaves and the wild type control. Metabolites were determined in samples harvested from leaves of four weeks old plants in the middle of the light or dark, respectively. Data are normalized to the mean response calculated for the wild type (individual wild-type values were normalized in the same way). Values presented are mean ± standard error of six individual plants. Values in bold denote significant differences that were determined by the t test (P < 0.05). Dark

Light

Wild type Amino acids Alanine ␤-Alanine Arginine Asparagine Aspartic acid Glutamic acid Glutamine Glycine Isoleucine Lysine Phenylalanine Proline Serine Threonine Tryptophan Tyrosine Valine Organic acids Pyruvic acid Aconitic acid Citric acid Succinic acid Fumaric acid Malic acid Maleic acid Lactic acid Glyceric acid Shikimic acid Ascorbic acid Dehydroascorbic acid Sugars Arabinose Fructose Glucose Sucrose Isomaltose Maltose Fucose Raffinose Trehalose Others Glucose-6-phosphate Fructose-6-phosphate Fructose-1.6-diphosphate Phosphoric acid 4-amino-butyric acid Benzoic acid Galactinol Glycerol Glycerol-3-phosphate Myo-inositol Mannitol Putrescine Spermidine Urea Nicotinic acid Nonanoic acid Octadecanoic acid

HIP1-3

Wild type

HIP1.3

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.15 0.07 0.05 0.15 0.04 0.04 0.09 0.16 0.19 0.12 0.09 0.16 0.07 0.08 0.06 0.11 0.13

0.75 1.15 1.59 1.60 1.02 1.04 1.48 2.78 1.70 1.25 1.40 1.79 1.13 1.65 1.77 1.12 1.47

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.08 0.16 0.27 0.35 0.08 0.07 0.25 0.70 0.26 0.09 0.15 0.41 0.14 0.15 0.13 0.07 0.19

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.127 0.100 0.095 0.191 0.120 0.043 0.088 0.264 0.133 0.127 0.094 0.202 0.056 0.130 0.174 0.106 0.203

1.26 0.79 0.57 0.57 0.80 0.88 0.66 0.80 1.02 0.74 0.77 0.73 0.86 1.32 0.90 0.67 1.03

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.16 0.05 0.05 0.10 0.04 0.04 0.10 0.20 0.19 0.07 0.06 0.09 0.06 0.11 0.06 0.07 0.23

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ±

0.05 0.05 0.04 0.11 0.05 0.05 0.05 0.13 0.16 0.05 0.19 0.08

1.24 1.05 0.98 1.79 0.93 0.96 0.98 0.79 0.50 1.19 1.87 1.32

± ± ± ± ± ± ± ± ± ± ± ±

0.09 0.07 0.02 0.31 0.02 0.02 0.02 0.06 0.07 0.11 0.29 0.10

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ±

0.209 0.216 0.056 0.127 0.060 0.068 0.039 0.098 0.077 0.138 0.320 0.052

0.85 0.67 0.81 0.57 0.85 0.82 0.90 0.90 0.58 0.86 0.25 0.68

± ± ± ± ± ± ± ± ± ± ± ±

0.17 0.06 0.03 0.01 0.06 0.02 0.05 0.15 0.04 0.05 0.03 0.05

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ±

0.07 0.28 0.25 0.05 0.14 0.05 0.06 0.14 0.06

1.21 2.26 2.28 0.94 1.38 1.27 1.24 1.64 1.11

± ± ± ± ± ± ± ± ±

0.08 0.37 0.17 0.02 0.12 0.13 0.09 0.21 0.09

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ±

0.161 0.175 0.155 0.058 0.222 0.195 0.149 0.160 0.170

0.90 0.83 0.89 0.84 0.56 0.81 0.78 0.65 0.83

± ± ± ± ± ± ± ± ±

0.05 0.04 0.04 0.04 0.05 0.05 0.03 0.07 0.04

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.05 0.05 0.17 0.09 0.14 0.08 0.12 0.21 0.08 0.04 0.25 0.07 0.10 0.14 0.09 0.10 0.11

0.88 0.93 2.60 1.15 1.50 0.91 1.53 0.77 1.11 1.11 0.83 0.91 0.98 1.06 0.98 1.18 1.16

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.08 0.10 0.38 0.19 0.22 0.09 0.20 0.10 0.03 0.06 0.11 0.07 0.10 0.13 0.10 0.14 0.05

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.125 0.161 0.246 0.223 0.157 0.156 0.142 0.155 0.111 0.048 0.247 0.101 0.188 0.133 0.114 0.124 0.131

0.85 0.88 0.67 0.82 0.86 0.90 0.52 0.90 0.80 0.80 0.68 0.63 0.54 0.72 0.71 0.72 0.97

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.08 0.09 0.08 0.04 0.11 0.12 0.09 0.11 0.07 0.04 0.08 0.06 0.06 0.05 0.06 0.05 0.12

varies when the leaves were harvested during the light or dark period (Table 2). Depending on the external conditions, the intracellular carbon fluxes are greatly altered. As the metabolic profiles of HIP1.3deficient mutant displays a major phenotype both in darkness and in the light it seems that HIP1.3 is permanently involved in the central carbon metabolism. However, the phenotype of the mutant

is qualitatively different in illuminated and darkened leaves. This might indicate that the biochemical function of HIP1.3 largely varies depending on the metabolic state of the leaf cells. As revealed by immuno blotting using buffer-soluble leaf proteins, the cellular content of HIP1.3 does not noticeably vary during the light and dark period (data not shown). Preliminary data indicate that leaves from the HIP1.3 deficient mutant that had been harvested during the light period differ

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largely from the wild type control in some galactose-containing oligosaccharides (such as melibiose; data not shown). These data are consistent with the assumption that HIP1.3 is indeed involved in the cytosolic carbohydrate metabolism. It should, however, be noted that the current knowledge on CBMs is far from being complete (Malik et al., 2010; Mello et al., 2010) and, therefore, the actual binding sites within the heteroglycans used by HIP1.3 cannot be predicted. Furthermore, binding of HIP1.3 to the cytosolic glycans suggests but does not necessarily imply that the protein exerts any catalytic activity on the very same molecule (see above). Finally, sequence-based predictions on the biochemical function of HIP1.3 are ambiguous and, therefore, require confirmation by empirical studies. Recently, data have been published that are related to the same product of gene At1g09340 but lead to conclusions largely deviating from those presented in this communication (Hassidim et al., 2007). These authors performed a data base search for putatively RNA binding proteins and, by doing so, they identified six genes one of these being At1g09340. As the transcript levels of this gene exhibit some diurnal fluctuations, it has been hypothesized that the product of gene At1g09340 is involved in the circadian control. Furthermore, this gene product exhibits some similarity to a plastidial endoribonuclease, CSP41a from Nicotiana tabaccum L. (Bollenbach et al., 2003) and, therefore, the same subcellular location and similar biochemical functions have been inferred. Because of these similarities, the gene has been designated as CHLOROPLAST RNA BINDING (CRB; Hassidim et al., 2007). However, none of the predictions has empirically been proven. CRB-deficient mutants grown in liquid culture in the presence of sucrose exhibited a reduction in size and structural alterations in most of the chloroplasts. These phenotypical features concur with those observed in soilgrown plants (Fig. 6a). However, the immunocytochemical analyses (Fig. 5b–d) clearly indicate that the target protein is located in the cytosol of mesophyll cells. However, within the cytosol the protein is unevenly distributed and, to some extent, is associated with the chloroplast envelope membranes. These data are fully consistent with the sequence-based predictions of the absence of a transitpeptide using the targetP and the chloroP programs (Emanuelsson et al., 2000, 1999). Both programs failed to strongly support any plastidial location of HIP1.3. In fact, according to targetP, HIP1.3 is more likely to be located in mitochondria or the endoplasmic reticulum as compared chloroplasts but, in any case, prediction is very weak. Furthermore, we were unable, despite some efforts, to experimentally demonstrate any RNA-binding activity of HIP1.3 (data not shown). By contrast, interaction with cytosolic heteroglycans was strong and easily demonstrable (Figs. 1–3). The product of gene At1g09340 does not exhibit noticeable changes throughout the light–dark cycle and, therefore, it is difficult to see that it directly involved in a circadian control. For all these reasons, it seems to us that the designation of the target protein as HIP1.3 is more appropriate. Acknowledgments Financial support by the Deutsche Forschungsgemeinschaft (SFB 429 ‘Molecular Physiology, Energetics, and Regulation of Primary Metabolism in Plants’ TP A11 ‘Regulation of respiration in plants’ and TP B2 ‘Structural and functional analysis of cytosolic highmolecular weight heteroglycans in higher plants’) is gratefully acknowledged. The authors thank Ms. Carola Kuhn (University of Potsdam, Plant Physiology) for performing the immunocytochemical studies. References Bollenbach TJ, Tatman DA, Stern DB. CSP41a, a multifunctional RNA-binding protein, initiates mRNA turnover in tobacco chloroplasts. Plant J 2003;36:842–52.

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