Research ArticleBIOCHEMICAL PROCESSES AND MACROMOLECULAR STRUCTURES
Open Access

Quantitative Proteomics of Seed Filling in Castor: Comparison with Soybean and Rapeseed Reveals Differences between Photosynthetic and Nonphotosynthetic Seed Metabolism

Norma L. Houston, Martin Hajduch, Jay J. Thelen

Published October 2009. DOI: https://doi.org/10.1104/pp.109.141622

Abstract

Seed maturation or seed filling is a phase of development that plays a major role in the storage reserve composition of a seed. In many plant seeds photosynthesis plays a major role in this process, although oilseeds, such as castor (Ricinus communis), are capable of accumulating oil without the benefit of photophosphorylation to augment energy demands. To characterize seed filling in castor, a systematic quantitative proteomics study was performed. Two-dimensional gel electrophoresis was used to resolve and quantify Cy-dye-labeled proteins expressed at 2, 3, 4, 5, and 6 weeks after flowering in biological triplicate. Expression profiles for 660 protein spot groups were established, and of these, 522 proteins were confidently identified by liquid chromatography-tandem mass spectrometry by mining against the castor genome. Identified proteins were classified according to function, and the most abundant groups of proteins were involved in protein destination and storage (34%), energy (19%), and metabolism (15%). Carbon assimilatory pathways in castor were compared with previous studies of photosynthetic oilseeds, soybean (Glycine max) and rapeseed (Brassica napus). These comparisons revealed differences in abundance and number of protein isoforms at numerous steps in glycolysis. One such difference was the number of enolase isoforms and their sum abundance; castor had approximately six times as many isoforms as soy and rapeseed. Furthermore, Rubisco was 11-fold less prominent in castor compared to rapeseed. These and other differences suggest some aspects of carbon flow, carbon recapture, as well as ATP and NADPH production in castor differs from photosynthetic oilseeds.

Castor (Ricinus communis) is a model heterotrophic oilseed that contains up to 60% fatty acids compared to approximately 20% and 40% in autotrophic oilseeds soybean (Glycine max) and rapeseed (Brassica napus), respectively (Weiss, 2000). Castor oil is a major component of many industrial lubricants and is currently of special interest, as it is being considered for biofuel use (Baldwin and Cossar, 2008; Dyer et al., 2008; Scholz and da Silva, 2008). With its high fatty acid content and recently sequenced genome, castor is a model plant for studying carbon assimilation in nonphotosynthetic oilseeds.

In oilseeds, fatty acid reserves are synthesized mainly during the seed-filling phase of seed development (Norton and Harris, 1975; Ruuska et al., 2002). Seed filling is a phase defined by morphological, cellular, and metabolic changes in the endosperm and embryo that coincide with rapidly increasing storage reserves, such as fatty acids and protein (Norton and Harris, 1975; Ruuska et al., 2002). Metabolic hallmarks of the seed-filling phase of castor seed development have been reported (Simcox et al., 1977; Greenwood and Bewley, 1981; Weiss, 2000). During development, seed length increases from 6 to 14 mm, while the seed coat color changes from white to red to speckled (Greenwood and Bewley, 1981). Fatty acid synthesis (FAS) begins in reddish-white seeds at approximately stage IV (3 weeks after flowering [WAF] for this study) and increases rapidly until stage VII (6 WAF) when the seed is speckled (Weiss, 2000). During this period, approximately 75% of fatty acid reserves are produced (Weiss, 2000).

De novo FAS requires carbon (acetyl-CoA), energy (ATP), and reducing equivalents (NADH/NADPH). In photosynthetic seeds, light energy can be used to produce ATP and NADPH within the plastid (Allen et al., 2009). In the plastids of heterotrophic seed, the origin of these components is unclear. Unlike ATP and NADH/NADPH, acetyl-CoA is not readily shuttled across the plastid membrane, where de novo FAS occurs (Bender-Machado et al., 2004). Therefore, precursors for acetyl-CoA synthesis must be produced in the plastid or imported from the cytosol. For example, Glc-6-P, pyruvate, and malate, may be imported via transporters or produced in the plastid (Smith et al., 1992; Eastmond et al., 1997; Pleite et al., 2005). When labeled carbon sources were supplied to isolated plastids from developing sunflower (Helianthus annuus) and castor seed, malate produced the highest rate of FAS (Smith et al., 1992; Eastmond et al., 1997; Lara-Núñez and Rodríguez-Sotres, 2004; Pleite et al., 2005). However, Alonso et al. (2007) suggested that malate only produces 5% to 9% of the carbon necessary for FAS in sunflower. Using metabolic flux analyses, they concluded that 91% to 95% of the carbon required for FAS was supplied by triose-Ps produced from hexoses imported into the plastid. The origin of energy and reducing equivalents needed for FAS also remains under investigation for nonphotosynthetic seeds. Studies conducted on heterotrophic plastids suggest two pathways are capable of providing adequate levels of reducing equivalents. One such pathway is the malate pathway described previously. Pleite et al. (2005) suggest that NADP-malic enzyme (ME) and the pyruvate dehydrogenase complex produce adequate levels of acetyl-CoA and reducing equivalents to support FAS in isolated plastids from sunflower embryos. Another pathway is the oxidative pentose-P pathway (OPPP), where three molecules of Glc-6-P enter the pathway and are oxidized to three molecules of ribulose-5-P and CO2 while forming six NADPH molecules (Kruger and von Schaewen, 2003). Alonso et al. (2007) showed that OPPP produces the majority of reducing equivalents for FAS in sunflower, compared to <4% produced by ME.

Proteomics can be a useful approach to study the integrated relationship between protein and oil composition in seed (Chen et al., 2009) as well as posttranscriptional regulatory mechanisms that would otherwise elude transcript, metabolite, or flux determinations. With continuing advances in global protein analyses, there have been several proteomic studies of seed filling in oilseeds, including soybean (Hajduch et al., 2005; Agrawal et al., 2008), rapeseed (Hajduch et al., 2006), and Medicago truncatula (Gallardo et al., 2003, 2007). From M. truncatula, Gallardo et al. (2003, 2007) identified a total of 224 proteins using two-dimensional gel electrophoresis (2-DGE) followed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) and matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF) during seed filling. Hajduch et al. (2006) used 2-DGE followed by LC-MS/MS and matrix-assisted laser desorption ionization time-of-flight mass spectrometry to identify 517 proteins during the seed-filling phase of rapeseed. These data were compared to 675 soybean proteins identified by 2-DGE and semicontinuous multidimensional protein identification technology (Sec-MudPIT) followed by LC-MS/MS (Hajduch et al., 2005; Agrawal et al., 2008). This comparative analysis showed increases in the number of rapeseed proteins involved in FAS (80% increase) and carbon assimilation proteins (48%) that lead to FAS (Agrawal et al., 2008). Results generated by these proteomic analyses and their comparisons provide further insight about metabolic pathway regulation during the seed-filling phase of photosynthetic oilseeds.

To characterize seed filling in a nonphotosynthetic seed, we report a quantitative proteomic analysis of castor seeds. We have examined the accumulation of proteins at five developmental stages of seed filling and established comprehensive whole-seed reference maps. From these gels, 660 spots of interest were detected, profiled, and analyzed by LC-MS/MS. Of these, 522 proteins were identified and sorted into 14 functional classes. These data were used to predict and compare the carbon assimilatory pathways in castor with those from soybean and rapeseed. Several differences in abundance and number of protein isoforms at numerous steps in these pathways were observed.

RESULTS

Characterization of Early Seed-Filling Phase in Developing Castor Seed

Castor seed harvesting was initiated at 2 WAF and continued at 7-d intervals until 6 WAF (Fig. 1A ). Each stage was characterized by seed coat color and seed length. Harvested castor seeds appeared white in color at 2 WAF, red at 3 WAF, and speckled brownish-white from 4 to 6 WAF. The seed coats were never green, consistent with the heterotrophic properties of castor seed. Seed length from 2 to 6 WAF ranged from 8 to 15 mm, in agreement with the range reported (5–16 mm) by Weiss (2000). Both color changes and seed length correspond to previously reports by Greenwood and Bewley (1981) and Weiss (2000). Based on these morphological hallmarks, each stage, 2 to 6 WAF, corresponded to previously described morphological stages II/III, IV, V, VI, and VII, respectively, noted by Greenwood and Bewley (1981).

Figure 1.
Figure 1.

Characterization of castor seed development during early seed-filling phase. A, Castor seeds at five experimental stages of development. Whole castor seeds were harvested at exactly 2, 3, 4, 5, and 6 WAF (left to right). Harvested seed corresponded to morphological stages II/III, IV, V, VI, and VII reported by Greenwood and Bewley (1981). B, Average fresh and dry seed mass at each experimental stage. Seed harvested at each experimental stage were weighed before and after drying. Values are the average of three biological replicates. sds are shown. C, Percentage of seed fresh mass. Percentages were calculated using the average fresh and dry masses. Average dry mass was subtracted from the average fresh mass to determine water content. Percentages of average dry mass and water content equal 100% of the average fresh mass. D and E, Total fatty acid and protein content of castor seed. Total fatty acids and protein are noted in milligrams/seed. Values are the average of three biological replicates. sds are shown. [See online article for color version of this figure.]

Fresh and dry seed masses and fatty acid and protein content were measured in all five experimental stages (Fig. 1, B–E). Both fresh and dry seed mass increased throughout seed development and were generally parallel in trend, suggesting the seed were in a predesiccation phase (Kermode and Bewley, 1985). These results also were reflected in the difference between the average fresh and dry masses, or water content, which decreased approximately 50% from 2 to 6 WAF (Fig. 1C). While the water content decreased, the total fatty acid content increased 50-fold from 2 to 6 WAF (Fig. 1, D and E). Both fatty acid and protein levels increased sharply between 5 and 6 WAF. Therefore, we chose to focus our study on the 2 through 6 WAF stages, which spanned the early to mid seed-filling phase. During this period of development, castor seed rapidly increased in protein and fatty acid content, but desiccation was not complete.

A Two-Step 2-DGE Approach Was Necessary to Produce High-Resolution Reference Maps for Castor Seed

Castor proteins were isolated from each experimental seed stage in biological triplicate for separation by 2-DGE. A two-step approach was employed to produce two-dimensional gels with minimal spot overlap and gel distortions. First, castor protein extracts were labeled with N-hydroxysuccinimide-activated Cy5 (NHS-Cy5), a Lys reactive fluorophore. Cyanine dyes have been used in two-dimensional difference gel electrophoresis to detect quantitative differences between two protein samples (Ünlü et al., 1997; Tonge et al., 2001). In this study, the advantage of using an NHS-activated Cy-dye was its specificity for Lys, an amino acid that is rare in storage proteins. Figure 2 shows the comparison of NHS-Cy5 labeled and Coomassie Brilliant Blue-stained proteins separated by 2-DGE. In Figure 2A, there are few instances of spot overlap and gel distortion despite the preponderance of seed storage proteins when compared to Coomassie Brilliant Blue-stained gel (Fig. 2B). NHS-Cy5-labeled proteins were resolved for analytical spot quantitation by both wide (pH 3–10) and medium (pH 4–7) range immobilized pH gradient (IPG) strips (Supplemental Fig. S1). The use of medium-range IPG strips enhanced spot separation in a dense area (pI 4–7) of the protein map.

Figure 2.
Figure 2.

2-DGE of Cy5-labeled and unlabeled proteins. Total protein was isolated from castor seeds harvested at 4 WAF. A, Protein extracts (0.05 mg) were labeled with fluorescent dye, Cy5, fractionated by 2-DGE, and detected using fluorescence imaging. B, Unlabeled protein extracts (1 mg) were fractionated by 2-DGE and stained with colloidal Coomassie Brilliant Blue. Migrating positions of isoelectric point (pI) and molecular mass (kD) markers are shown. Circles highlight corresponding regions.

LC-MS/MS Analyses Yielded 522 Identified Proteins

Gel images were scanned and analyzed with ImageMaster to detect, quantify, and match spots. Relative volume, defined as the ratio of each spot volume per total spot volume, was calculated for each spot (Supplemental Table S1). To merge data from medium- and wide-range gels, spot volumes were adjusted with a correction constant (Hajduch et al., 2005). Corrected relative volumes were averaged, graphed, and displayed as protein expression profiles. Spots from each gel were matched within biological replicates and between developmental stages (Fig. 3A ). To be considered for identification, spots had to meet two criteria: (1) present in at least two of five developmental stages and (2) present in three biological replicates of each developmental stage. Matched spots that met these criteria were termed spot groups. A total of 660 spot groups met these criteria. These spots were excised from colloidal Coomassie Brilliant Blue-stained reference gels to ensure sufficient protein for mass spectrometry and to prevent any misassignments caused by Mr shifting of labeled proteins (Ünlü et al., 1997). After excision, spots were trypsin digested, analyzed by LC-MS/MS, and searched against The Institute for Genomic Research (TIGR) castor translated genome using SEQUEST (Fig. 3B), resulting in identification of 522 (79%) out of 660 spot groups. Of these 522 proteins, 303 were nonredundant. Protein assignment information and expression profiles of these spot groups were deposited in the Oilseed Proteomics Web database (http://oilseedproteomics.missouri.edu/).

Figure 3.
Figure 3.

Summary of experimental approach. A, Expression profile workflow. Protein extracts from each experimental stage were fractionated by 2-DGE in biological triplicate. Gels were imaged and analyzed to detect, quantify, and match spots. Spots represented in all three gels and at least two experimental stages were matched and labeled as spot groups. Average relative volumes, sds, and expression profiles were developed for the 660 spot groups. B, Protein identification workflow. Six hundred and sixty spots were excised from colloidal Coomassie Brilliant Blue reference gels. Each protein spot was digested, and the peptides were analyzed by LC-MS/MS. Spectra were searched against the castor TIGR database using SEQUEST. All protein assignments and expression profiles were deposited in the oilseeds proteomics Web site.

Identified castor proteins were grouped into (Supplemental Table S2) functional classifications according to Bevan et al. (1998) with modifications by Hajduch et al. (2006) for seed tissue. Classified castor proteins were compared to parallel 2-DGE proteomic studies of soybean and rapeseed (Hajduch et al., 2006; Agrawal et al., 2008; Fig. 4 ; Supplemental Table S3). There were striking similarities in the distribution of proteins from the three oilseeds. Functional classes with the highest percentage of proteins were destination/storage, energy, and primary metabolism for all three oilseeds. As expected, soybean, a high protein oilseed, had the highest percentage (44%) of protein in the destination/storage class followed by castor (34%) and rapeseed (25%). Rapeseed (23%) had the highest percentage of proteins involved in energy production followed by castor (19%) and soybean (11%). All three oilseeds showed similar percentages of proteins in primary metabolism (castor 15%, rapeseed 17%, and soybean 16%).

Figure 4.
Figure 4.

Functional classification of identified proteins from castor, soybean, and rapeseed. Castor proteins were identified and grouped into 14 functional classes as described by Bevan et al. (1998) and modified by Hajduch et al. (2006). These data were compared with parallel studies of seed filling in soybean and rapeseed (Hajduch et al., 2006; Agrawal et al., 2008). The histogram represents percentage of identified proteins in each functional class. Numbers in parentheses correspond to functional classifications in Supplemental Table S3.

To understand the progression of metabolic activities during the early seed-filling phase of castor seed development, composite expression profiles were created for each functional subclass (Hajduch et al., 2005). Pooled relative volumes were calculated by summing relative volumes of identified proteins in each subclass and expression profiles graphed. For simplicity, only those subclasses with 10 or more proteins were included in this analysis. As a result, 79% of identified proteins were grouped into 12 subclasses (Fig. 5 ; Supplemental Table S4). Two of the three classes with the largest relative abundance were in the protein destination/storage functional class. The storage protein subclass had the largest relative abundance and showed an increasing expression pattern beginning at 3 WAF. The glycolysis subclass, a member of the energy functional class, was the second largest class of proteins and showed a steady decrease in expression. The third largest class was folding and stability, which showed an expression profile that increased at 3 WAF but declined steadily thereafter. Overall, the expression patterns of the 12 subclasses were categorized into three groups based on their expression pattern during seed development (Fig. 5). The first group included nine of the 12 subclasses and was composed of subclasses that are expressed mainly during the early stages of seed development. Group two, which only included the detoxification subclass, was expressed generally in the middle stages of development. The expression pattern of this group showed little change in relative abundance during development. Finally, group three was expressed mainly in the later stages of the experimental phase and included defense-related proteins and storage protein subclasses.

Figure 5.
Figure 5.

Functional subclasses grouped according to expression pattern during castor seed development. Composite expressions of protein belonging to functional subclasses were grouped according to their similarity (Hajduch et al., 2005). Only subclasses with 10 or more proteins are shown. Number of proteins (right) and maximum relative volumes (left) are shown above the expression profile. Numbers in parentheses correspond to functional subclassifications in Supplemental Table S4.

Of the identified proteins, the 20 spot groups with the largest overall relative abundance are listed in Table I . To calculate the overall relative abundance throughout seed filling, the average relative volumes from each experimental stage of a single protein spot were summed. Because of their well-documented prominence in seeds, storage proteins were excluded from this analysis. Interestingly, eight out of the 20 proteins were energy related and specifically involved in glycolysis. These proteins include glyceraldehyde 3-P dehydrogenase (GAPDH), enolase, Fru-bisP aldolase (FBA), and phosphoglycerate kinase (PGK). Six of the 20 proteins were involved in protein folding and stability and were annotated as either protein disulfide isomerase (PDI) or heat shock proteins. A PDI protein (spot 185) was the most abundant nonstorage protein expressed during castor seed filling. High levels of PDI and heat shock proteins may assist in the folding of large amount of storage proteins (Houston et al., 2005). Together, these 20 proteins represent 11% to 20% of the pooled relative volume during castor seed filling.

View this table:
Table I.

Most abundant proteins expressed during seed filling in castor

DISCUSSION

Quantitative Proteomics Suggest Differences in Carbon Assimilation between Photosynthetic and Nonphotosynthetic Oilseeds

To compare carbon assimilation in castor to photosynthetic oilseeds, soybean and rapeseed, metabolic enzymes were mapped to glycolysis and C4/C3 organic acid metabolism leading to FAS (Hajduch et al., 2005; Agrawal et al., 2008). In both figures (Figs. 6 and 7 ), enzymes are mapped to show the production of phosphoenolpyruvate (PEP) in the cytosol and plastid, but from PEP production there are two potential pathways. First, in the flow of C3 carbon, pyruvate kinase (PK) converts PEP into pyruvate in the cytosol or plastid; second, in the flow of C4 carbon, phosphoenolpyruvate carboxylase (PEPC) carboxylates PEP to produce oxaloacetate (OAA) and then malate via malate dehydrogenase (MDH). Figures 6 and 7 also show the comparison of expression patterns, maximum relative abundance, number of isoforms, and subcellular localization of these identified proteins (Hajduch et al., 2005; Agrawal et al., 2008). These comparisons reveal differences in abundance and number of protein isoforms at many of these steps. The differences suggest malate may have a more prominent role in intermediary metabolism in castor than in photosynthetic oilseeds soybean and rape.

Figure 6.
Figure 6.

Characterization of carbon assimilation during seed filling in oilseeds. Proteins involved in sugar breakdown that lead to carbon sources for biosynthetic pathways are displayed. Expression profiles show the relative abundance for each enzyme at each stage of the early seed-filling phase. C (castor), S (soybean), or R (rapeseed) above the expression profile represents the origin of the protein (Hajduch et al., 2006; Agrawal et al., 2008). The number above the graph shows the number of identified proteins. The number to the left of the graph shows the maximum value (y axis) of the relative volume. Solid lines represent proteins identified in parallel proteomic studies of castor, soybean, or rapeseed, while unidentified proteins are represented by dashed lines. Checked squares indicate identified proteins with no expression profiles due to low volumes or presence in less than three stages. Brackets indicate the number of proteins identified by Sec-MudPIT. Gray boxes highlight differences in number (≥2) or relative abundance (≥2). Abbreviations not defined in the text: SuSy, Suc synthase; UGP, UDP-Glc pyrophosphorylase; PGM, phosphoglucomutase; PGI, phosphoglucose isomerase; FK, Fru kinase; PFK, phosphofructose kinase; TPI, triose-P isomerase; PDC, pyruvate dehydrogenase complex.

Figure 7.
Figure 7.

Characterization of malate synthesis pathways. Expression profiles show the relative abundance (left), species of origin (C, castor; S, soybean; R, rapeseed), and number of identified proteins (top). Solid lines represent proteins identified in parallel proteomic studies of castor, soybean, or rapeseed, while unidentified proteins are represented by dashed lines. Brackets indicate proteins identified by immunoblot analysis. The proteins shown are listed as follows: PEPC, MDH, ME, PK, and PDC (pyruvate dehydrogenase complex).

Five Glycolytic Proteins Showed an Increased Number of Isoforms and Abundance When Compared to Soybean and Rapeseed

Glycolysis is capable of producing, either directly or indirectly, necessary carbon sources, reducing equivalents, and energy required for FAS (Plaxton and Podestá, 2006). For instance, Schwender et al. (2003), Alonso et al. (2007), and Iyer et al. (2008) suggest that FAS is primarily supported by glycolysis in B. napus embryo cultures, sunflower embryos, and soybean cotyledons. Classical glycolysis (i.e. via hexokinase, ATP-phosphofructose kinase, GAPDH, and PK) has a net yield of two pyruvate, two ATP, and two NADH for every Glc molecule that enters the pathway (for review, see Plaxton and Podestá, 2006). Also, glycolytic intermediates and products, such as Glc-6-P, PEP, and pyruvate, may indirectly lead to similar components through the tricarboxylic acid (TCA) cycle, malate synthesis, or the OPPP. For example, PEP can be transported across plastid membranes and then used by PK to produce ATP and pyruvate (Ruuska et al., 2002). Therefore, we will highlight selected glycolytic enzymatic steps between triose-P and pyruvate to investigate their contribution to FAS. Because glycolysis in plants occurs in both the cytosol and plastid, subscripts of c (cytosol) and p (plastid) are used to indicate predicted subcellular localization when necessary.

The first glycolytic enzyme that was differentially expressed in castor compared to soybean and rapeseed is cytosolic FBA. This enzyme was 30% higher in castor than rapeseed, while soybean expression levels were undetectable. FBA catalyzes an aldol cleavage of Fru-1,6-bisP to triose-Ps, dihydroxyacetone phosphate, and glyceraldehyde 3-phosphate (GAP). Alonso et al. (2007) suggested that triose-Ps are the major carbon source for FAS in sunflower embryos, another nonphotosynthetic oilseed. The increase in FBAc for castor may be necessary to produce large amounts of triose-Ps. Two of the eight FBAc isoforms (spots 344 and 356) were in the top 10 overall abundant proteins. By contrast, four FBAp isoforms were identified. In green leaves of maize (Zea mays), wheat (Triticum aestivum), spinach (Spinacia oleracea), and pea (Pisum sativum), most of the FBA activity (approximately 90%) is chloroplastic (Schnarrenberger and Krüger, 1986; Pelzer-Reith et al., 1993), and a small decrease in FBAp activity in potato (Solanum tuberosum) leaves resulted in the inhibition of photosynthesis and starch synthesis (Haake et al., 1998). However, in germinating castor seed, FBA activity is mostly cytosolic, representing two-thirds of the activity (Moorhead et al., 1994). In developing castor seed, FBAc is 5-fold more abundant than FBAp in contrast to what was observed in green leaves (Haake et al., 1998). Alternatively, Simcox et al. (1977) showed higher FBAp activity in developing castor seed, although the plastid fractionation was contaminated with other cytosolic enzymes.

GAPDH converts GAP to glycerate-1,3-bisP. There are four types of GAPDH in plants: (1) cytosolic, NAD-dependent GAPC; (2) cytosolic, nonphosphorylating GAPDH; (3) chloroplastic, NADP-specific GAPA/B (Cerff and Chambers, 1979); and (4) heterotrophic plastids, GAPCp (Petersen et al., 2003; Rius et al., 2008). Of these, GAPC is the active isoform in cytosolic glycolysis (Valverde et al., 2005; Hajirezaei et al., 2006). In castor, 12 GAPC isoforms were identified, and three of these proteins (spots 383, 385, and 387) were quantified as the most abundant proteins expressed in developing castor. Castor had >2 times the isoforms and abundance of soybean and rapeseed GAPC. The pattern of GAPC expression was similar to FBA. NP-GAPDH was identified as part of a bypass of GAPC that converts GAP to 3-phosphoglycerate (Valverde et al., 2005; Supplemental Fig. S2). Only one NP-GAPDH isoform (spot 230) was identified. The low number of isoforms and relative abundance suggest that this may not play a major activity in castor seeds. Finally, two GAPCp isoforms (spots 369 and 372) were identified, but expression was 10-fold lower than GAPC in castor as well as soybean and rapeseed. The GAPCp expression profile during seed filling was similar to that of GAPC in all three oilseeds. All of the identified GAPDH isoforms are capable of producing reducing equivalents. The high level of GAPDH expression, especially GAPC, may be related to an increased need for reducing equivalents or glycolytic intermediates.

Two other glycolytic activities that were more prominent in castor compared to soybean and rapeseed are PGK and 2,3-bisphosphoglycerate-independent phosphoglycerate mutase (iPGAM). First, PGK converts glycerate-1,3-bisP to 3-phosphoglycerate while producing ATP; it is one of only two glycolytic steps that produce ATP. One of the four PGKc 2-DGE forms (spot 1162) represents about half of the total relative abundance. Interestingly, both PGK and iPGAM levels were about 5- to 6-fold lower than the previous two steps catalyzed by FBA and GAPDH. Castor has the highest number of iPGAM isoforms and relative abundance compared to soybean and rapeseed. Expression patterns of castor and soybean are similar, but the most obvious pattern is the decreasing pattern continued from the previous glycolytic enzymes. Another interesting pattern is the decrease in PGK and iPGAM isoforms and volume in castor, soybean, and rapeseed when compared to the previous steps catalyzed by FBA and GAPDH. The conservation of this decrease may be of regulatory importance to control the flow of carbon or may reflect a higher Kcat for these two enzymes.

Miernyk and Dennis (1992) previously showed that enolase activity in castor seeds was highest during FAS and then decreased to undetectable levels. These results are reflected in this study. Enolase catalyzes the conversion of 2-PGA to PEP. Castor had six times as many enolase isoforms than soybean and rapeseed during the seed-filling phase. Two of these isoforms were classified as the most abundant proteins overall. Collectively, the enolase isoforms produced a relative abundance 2 to 3 times higher than soybean and rapeseed. There is a paucity of literature to explain this high expression, but a simple explanation is that it is necessary to accommodate increased flux of triose-P from both glycolysis and carbon dioxide recycling. This explanation is supported by similar expression patterns and increased numbers of isoforms and expression of four prior glycolytic steps. Alternatively, if PEPC is involved in CO2 sequestering, as previously suggested, PEP production will be at a premium (Rodríguez-Sotres et al., 2000). Significant rates of dark CO2 fixation from intact developing castor seeds are consistent with this suggestion (Benedict and Beevers, 1961).

In the final step of glycolysis, PK catalyzes the irreversible transfer of inorganic phosphate from PEP to ADP, yielding pyruvate and ATP. This step is considered to be a highly regulated glycolytic step as it is activated by pH changes, inhibited by Glu/Asp and ATP, and degraded after phosphorylation and ubiquitination (Podesta and Plaxton, 1991; Tang et al., 2003; Turner et al., 2005). Among the oilseeds compared, PKc isoforms were identified in soybean and castor, while the PKp was identified in rapeseed; however, in earlier studies, PKp activity and concentration was shown to coincide with fatty acid levels in developing castor seed (Plaxton, 1991; Negm et al., 1995). Castor and soybean expression patterns are very different. PKc expression in castor has a steady decreasing trend consistent with other glycolytic enzymes, while soybean shows a more anomalous trend, perhaps suggestive of more complex regulation.

Schwender et al. (2003) demonstrated that glycolysis supplies most carbon to FAS in rapeseed developing embryos in culture. Alonso et al. (2007) suggested that triose-Ps are the carbon sources for FAS in heterotrophic sunflower embryos, but they were unable to determine the source of triose-P production. In castor seed, the bulk of expressed glycolytic enzymes were cytosolic, and the five glycolytic steps between triose-Ps and PEP were prominent in both volume and number of isoforms. Hewezi et al. (2006) observed similar results in sunflower leaves and immature embryos using cDNA microarrays. This study showed a coordinated up-regulation of glycolytic genes (e.g. FBAc, GAPDHc, and PKp) in immature embryos compared to leaves (Hewezi et al., 2006). One specific similarity was the up-regulation of GAPDHc transcript in sunflower (a heterotrophic oilseed), while our proteomic analysis of castor revealed four and two times as many isoforms as soybean and rapeseed, respectively.

Enzymes Involved in C4 Intermediary Metabolism Are More Prominent in Castor Compared to Soybean and Rapeseed

Malate has been shown to support FAS in isolated castor and sunflower seed plastids and has been investigated as the major source of carbon skeletons and reducing power for FAS (Eastmond et al., 1997; Pleite et al., 2005). Figure 7 shows a diagram of malate synthesis while comparing the expression patterns, maximum relative abundance, number of isoforms, and subcellular localization of identified proteins in castor, soybean, and rapeseed (Hajduch et al., 2005; Agrawal et al., 2008). In the cytosol, the first step in malate production is performed by PEPC, which catalyzes the irreversible β-carboxylation of PEP to yield OAA and inorganic phosphate. PEPC is a ubiquitous protein in vascular plants and plays pivotal roles in replenishing the TCA cycle and fixing carbon dioxide in C4 and CAM plants (Chollet et al., 1996; Izui et al., 2004). In heterotrophic plants, PEPC has been postulated to provide carbon precursors for FAS through its role in malate synthesis and recycling of carbon dioxide emitted from the pyruvate dehydrogenase complex (Eastmond et al., 1997; Pleite et al., 2005; Uhrig et al., 2008). PEPC could also play a role in recycling CO2 from malic enzyme, as well as the decarboxylating reactions of the TCA cycle. Our proteomic analysis did not identify PEPC possibly due to its high Mr. However, Sangwan et al. (1992) have shown that the most significant increase in PEPC activity and concentration coincides with the increase of fatty acid accumulation in developing castor seeds.

The presence of MDH and ME suggests a contribution of malate synthesis to intermediary metabolism in castor seed. MDH followed by ME converts OAA to pyruvate in two reactions. MDH catalyzes the reversible conversion of OAA to malate (Goward and Nicholls, 1994). Among the oilseeds compared, castor has the lowest number of cytosolic MDH isoforms and overall relative volume. However, castor has the only MDH isoform identified from the plastid. Expression of the plastid isoform suggests a role for malate in the transfer of reduction equivalents from the plastid to the cytosol. Unlike MDH, ME was only identified in castor seed. In developing castor seed, plastid NADP-ME activity was shown to peak at later stages of seed development as lipid production increased (Shearer et al., 2004); however, cytosolic ME had the higher volume when compared to the plastid isoform and may also be important in supplying carbon and reducing equivalents for FAS in developing seed. Overall, it appears that the cytosolic pathway of malate synthesis is the major pathway in developing castor seed.

The First Three Proteins of OPPP Were Identified

The OPPP is also capable of being a major source of reducing equivalents for FAS (Schwender et al., 2003; Alonso et al., 2007). Reducing equivalents are produced in the first three steps of the OPPP by Glc-6-P dehydrogenase, gluconate-6-P lactonase, and gluconate-6-P dehydrogenase (Supplemental Fig. S3). Both cytosolic and plastidal isoforms of Glc-6-P dehydrogenase and gluconate-6-P dehydrogenase have been identified in photosynthetic and nonphotosynthetic tissues (Graeve et al., 1994; von Schaewen et al., 1995; Wendt et al., 2000; Knight et al., 2001; Krepinsky et al., 2001), but some studies indicate the castor plastid lacks a complete or efficient OPPP. For example, studies indicate that castor seed plastids do not contain Glc-6-P dehydrogenase, a key enzyme in the pentose-P pathway (Simcox et al., 1977; Nishimura and Beevers, 1979). Our analysis identified putative cytosolic and plastidal Glc-6-P dehydrogenases but only cytosolic gluconate-6-P lactonase and plastidal gluconate-6-P dehydrogenase. All identified Glc-6-P dehydrogenase proteins show decreasing expression pattern during seed filling; however, the number of identified proteins and their relative abundance was higher for the putative cytosolic isoforms. Both steps catalyzed by Glc-6-P and gluconate-6-P dehydrogenase produce NADPH and were identified in castor. Of the OPPP identified proteins, the putative plastidal gluconate-6-P dehydrogenase showed the largest relative abundance. The relative abundance of gluconate-6-P dehydrogenase was 3- and 6-fold higher than that of soybean and rapeseed.

CONCLUSION

This study provides a quantitative, proteomics analysis of seed filling in a nonphotosynthetic seed. The results of this study and comparison with parallel proteomic studies of photosynthetic oilseeds enhance our current knowledge of both seed filling and carbon metabolism in a nonphotosynthetic tissue. Using the data from our proteomic analysis, we mapped the activities to various carbon assimilatory pathways. The data show an increase in number and abundance of five glycolytic proteins in castor compared to soybean and rapeseed. We also demonstrate the presence of proteins for a complete cytosolic malate synthesis pathway that was absent in prior parallel studies of rapeseed and soybean. Our data corroborate previous studies of isolated leucoplasts, PEPC, and ME from developing castor seed, suggesting that cytosolic glycolysis and malate synthesis are likely contributors of both carbon and reducing equivalents for FAS.

MATERIALS AND METHODS

Plant Material and Growth Conditions

Castor (Ricinus communis) was grown in a greenhouse in Columbia, Missouri, with supplemental lighting (16-h-light/8-h-dark cycle, 26°C day/21°C night cycle). Plants were fertilized (15:30:15) once every 2 weeks. Castor flowers were tagged between 1 and 3 pm CST, and developing seed were collected precisely at 2, 3, 4, 5, and 6 WAF.

Protein Isolation, Cy-Dye Labeling, and 2-DGE

Total protein was isolated from developing seed and subjected to 2-DGE as described previously (Hajduch et al., 2005). Protein quantification was performed in triplicate using the Coomassie dye binding assay (Bio-Rad) against standard curves of chicken γ-globulin.

For Cy5 labeling, protein pellets were reconstituted in resuspension buffer (30 mm Tris-HCl, pH 8.5, 7 m urea, 2 m thiourea, and 4% [w/v] CHAPS) with vortex mixing for 30 min at room temperature followed by centrifugation for 15 min at 14,000g to remove insoluble material. Then, 50 μg of protein was taken and adjusted to final volume of 10 μL with resuspension buffer. One microliter of Cy-Dye (100 pmol) was added and incubated on ice for 30 min in the dark. The labeling reaction was terminated with 1 μL of 10 mm Lys on ice for an additional 10 min in the dark. For isoelectric focusing (IEF), 50 μg of protein was mixed with equal volume of 2× sample buffer (8 m urea, 130 mm dithiothreitol, and 4% [w/v] CHAPS), incubated 10 min on ice, mixed with 2.25 μL of IPG buffer (GE Healthcare), and adjusted to total volume of 450 μL with sample buffer.

For preparative, colloidal Coomassie Brilliant Blue-stained gels, protein pellets were resuspended in IEF sample extraction buffer (8 m urea, 2 m thiourea, 2% [w/v] CHAPS, 2% [v/v] Triton X-100, and 50 mm dithiothreitol) with vortex mixing as described above. For IEF, 1 mg of total protein was mixed with 2.25 μL of appropriate IPG buffer in a total volume of 450 μL of IEF extraction buffer.

Image Acquisition and Analysis

Fluorescent gels were scanned using a FLA-5000 laser scanner (FUJI Medical). Preparative Coomassie Brilliant Blue-stained gels were imaged by scanning densitometry (300 dpi, 16-bit grayscale pixel depth) as described previously (Hajduch et al., 2007). Digitized images were analyzed using ImageMaster 2-D Platinum software (version 5.0; GE Healthcare) in biological triplicate according to Hajduch et al. (2005). Protein abundance was expressed as relative volume according to the normalization method provided by ImageMaster software. Relative volumes were adjusted with correction constants to merge data from both 4 to 7 and 3 to 10 gels. Relative volumes were averaged and graphed.

Protein Identification by Mass Spectrometry

Proteins spots represented in all three biological and technical replicates were excised from corresponding preparative Coomassie Brilliant Blue-stained two-dimensional gel and trypsin digested as described previously (Hajduch et al., 2005). Mass spectral analysis of trypsin-digested protein samples were carried out on a linear ion trap tandem mass spectrometer (ProteomeX LTQ-XL; Thermo-Fisher) using liquid chromatography and nanospray ionization. The LC-MS/MS was operated according to Hajduch et al. (2006) for high-throughput protein identification.

Database Searching with Spectral Data and Uploading to the Oilseed Proteome Database

Tandem mass spectra were mined against the castor translated genome (June, 2007) using SEQUEST (Eng et al., 1994; Yates et al., 1995) as part of the BioWorks version 3.3 program. Search parameters were as follows: enzyme, trypsin; number of internal cleavage sites, 2; mass range, 400 to 1600; threshold, 500; minimum ion count, 35; peptide mass tolerance, 1.50 D; variable modifications, oxidation (M); static modification, carboxyamidomethylation (C). Matching peptides were filtered according to cross-correlation scores (XCorr at least 1.5, 2.0, and 2.5 for +1, +2, and +3 charged peptides, respectively) and peptide probability (0.05 or lower). For all protein assignments, a minimum of two unique peptides was required. All data from this investigation are available from the oilseed proteomics Web database (http://oilseedproteomics.missouri.edu). Programming for the Web database was performed as described previously (Hajduch et al., 2005). Data are viewable as active links from two-dimensional gels and as a protein identification table.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure S1. Two-dimensional reference maps of castor proteins isolated from seeds at 2, 3, 4, 5, and 6 WAF.

  • Supplemental Figure S2. Characterization of GAPDH reactions in castor seeds.

  • Supplemental Figure S3. Characterization of OPPP (first three steps).

  • Supplemental Table S1. Corrected relative volumes for spot groups.

  • Supplemental Table S2. Identified castor proteins listed according to functional class.

  • Supplemental Table S3. Percentage of identified proteins from castor, rapeseed, and soybean.

  • Supplemental Table S4. Pooled relative volumes of functional classes in castor seeds.

Acknowledgments

We are grateful to Ganesh Agrawal for technical assistance. We also thank Jianjiong Gao for uploading the castor data to the Proteomics of Oilseeds Web site.

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Jay J. Thelen (thelenj{at}missouri.edu).

  • www.plantphysiol.org/cgi/doi/10.1104/pp.109.141622

  • ↵1 This work was supported by the National Science Foundation Plant Genome Research Program Young Investigator Award (grant no. DBI–0332418 to J.J.T.).

  • ↵2 Present address: Institute of Plant Genetics and Biotechnology, Slovak Academy of Sciences, 95007 Nitra, Slovak Republic.

  • ↵[C] Some figures in this article are displayed in color online but in black and white in the print edition.

  • ↵[OA] Open access articles can be viewed online without a subscription.

  • ↵[W] The online version of this article contains Web-only data.

  • Received May 18, 2009.
  • Accepted August 11, 2009.
  • Published August 12, 2009.

LITERATURE CITED

  1. ↵
    Agrawal GK, Hajduch M, Graham K, Thelen JJ (2008) In-depth investigation of soybean seed-filling proteome and comparison with a parallel study of rapeseed. Plant Physiol 148: 504–518
    OpenUrlAbstract/FREE Full Text
  2. ↵
    Allen D, Ohlrogge JB, Shachar-Hill Y (2009) The role of light in soybean seed filling metabolism. Plant J 58: 220–234
    OpenUrlCrossRefPubMed
  3. ↵
    Alonso AP, Goffman FD, Ohlrogge JB, Shachar-Hill Y (2007) Carbon conversion efficiency and central metabolic fluxes in developing sunflower (Helianthus annuus L.) embryos. Plant J 52: 296–308
    OpenUrlCrossRefPubMed
  4. ↵
    Baldwin BS, Cossar RD (2008) Castor yield in response to planting date at four locations in the south-central United States. Ind Crops Prod 29: 316–319
    OpenUrlCrossRef
  5. ↵
    Bender-Machado L, Bäuerlein M, Carrari F, Schauer N, Lytovchenko A, Gibon Y, Kelly A, Loureiro M, Müller-Röber B, Willmitzer L, et al (2004) Expression of a yeast acetyl CoA hydrolase in the mitochondrion of tobacco plants inhibits growth and restricts photosynthesis. Plant Mol Biol 55: 645–662
    OpenUrlCrossRefPubMed
  6. ↵
    Benedict CR, Beevers H (1961) Formation of sucrose from malate in germinating castor beans. I. Conversion of malate to phosphoenol-pyruvate. Plant Physiol 36: 540–544
    OpenUrlFREE Full Text
  7. ↵
    Bevan M, Bancroft I, Bent E, Love K, Goodman H, Dean C, Bergkamp R, Dirkse W, Van Staveren M, Stiekema W, et al (1998) Analysis of 1.9 Mb of contiguous sequence from chromosome 4 of Arabidopsis thaliana. Nature 391: 485–488
    OpenUrlCrossRefPubMed
  8. ↵
    Cerff R, Chambers SE (1979) Subunit structure of higher plant glyceraldehyde-3-phosphate dehydrogenases (EC 1.2.1.12 and EC 1.2.1.13). J Biol Chem 254: 6094–6098
    OpenUrlAbstract/FREE Full Text
  9. ↵
    Chen M, Mooney BP, Hajduch M, Joshi T, Zhou M, Xu D, Thelen JJ (2009) System analysis of an Arabidopsis mutant altered in de novo fatty acid synthesis reveals diverse changes in seed composition and metabolism. Plant Physiol 150: 27–41
    OpenUrlAbstract/FREE Full Text
  10. ↵
    Chollet R, Vidal J, O'Leary MH (1996) Phosphoenolpyruvate carboxylase: a ubiquitous, highly regulated enzyme in plants. Annu Rev Plant Physiol Plant Mol Biol 47: 273–298
    OpenUrlCrossRef
  11. ↵
    Dyer JM, Stymne S, Green AG, Carlsson AS (2008) High-value oils from plants. Plant J 54: 640–655
    OpenUrlCrossRefPubMed
  12. ↵
    Eastmond PJ, Dennis DT, Rawsthorne S (1997) Evidence that a malate/inorganic phosphate exchange translocator imports carbon across the leucoplast envelope for fatty acid synthesis in developing castor seed endosperm. Plant Physiol 114: 851–856
    OpenUrlAbstract
  13. ↵
    Eng JK, McCormack AL, Yates JR (1994) An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J Am Soc Mass Spectrom 5: 976–989
    OpenUrlCrossRefPubMed
  14. ↵
    Gallardo K, Firnhaber C, Zuber H, Héricher D, Belghazi M, Henry C, Küster H, Thompson R (2007) A combined proteome and transcriptome analysis of developing Medicago truncatula seeds: evidence for metabolic specialization of maternal and filial tissues. Mol Cell Proteomics 6: 2165–2179
    OpenUrlAbstract/FREE Full Text
  15. ↵
    Gallardo K, Signor CL, Vandekerckhove J, Thompson RD, Burstin J (2003) Proteomics of Medicago truncatula seed development establishes the time frame of diverse metabolic processes related to reserve accumulation. Plant Physiol 133: 664–682
    OpenUrlAbstract/FREE Full Text
  16. ↵
    Goward CR, Nicholls DJ (1994) Malate dehydrogenase: a model for structure, evolution, and catalysis. Protein Sci 3: 1883–1888
    OpenUrlCrossRefPubMed
  17. ↵
    Graeve K, von Schaewen A, Scheibe R (1994) Purification characterisation and cDNA sequence of glucose-6-phosphate dehydrogenase from potato (Solanum tuberosum L.). Plant J 5: 535–561
    OpenUrlCrossRef
  18. ↵
    Greenwood JS, Bewley JD (1981) Seed development of Ricinus communis (castor bean) I. Descriptive morphology. Can J Bot 60: 1751–1760
    OpenUrlCrossRef
  19. ↵
    Haake V, Zrenner R, Sonnewald U, Stitt M (1998) A moderate decrease of plastid aldolase activity inhibits photosynthesis, alters the levels of sugars and starch, and inhibits growth of potato plants. Plant J 14: 147–157
    OpenUrlCrossRefPubMed
  20. ↵
    Hajduch M, Casteel JE, Hurrelmeyer KE, Song Z, Agrawal GK, Thelen JJ (2006) Proteomic analysis of seed filling in Brassica napus. Developmental characterization of metabolic isozymes using high-resolution two-dimensional gel electrophoresis. Plant Physiol 141: 32–46
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Hajduch M, Casteel JE, Tang S, Hearne LB, Knapp SK, Thelen JJ (2007) Proteomic analysis of near-isogenic sunflower varieties differing in seed oil traits. J Proteome Res 6: 3232–3241
    OpenUrlCrossRefPubMed
  22. ↵
    Hajduch M, Ganapathy A, Stein JW, Thelen JJ (2005) A systematic proteomic study of seed filling in soybean. Establishment of high-resolution two-dimensional reference maps, expression profiles, and an interactive proteome database. Plant Physiol 137: 1397–1419
    OpenUrlAbstract/FREE Full Text
  23. ↵
    Hajirezaei M-R, Biemelt S, Peisker M, Lytovchenko A, Fernie AR, Sonnewald U (2006) The influence of cytosolic phosphorylating glyceraldehyde 3-phosphate dehydrogenase (GAPC) on potato tuber metabolism. J Exp Bot 57: 2363–2377
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Hewezi T, Petitprez M, Gentzbittel L (2006) Primary metabolic pathways and signal transduction in sunflower (Helianthus annuus L.): comparison of transcriptional profiling in leaves and immature embryos using cDNA microarrays. Planta 223: 948–964
    OpenUrlCrossRefPubMed
  25. ↵
    Houston NL, Chuanzhu F, Xiang Q-Y, Schulze J-M, Jung R, Boston RS (2005) Phylogenetic analyses identify 10 classes of the protein disulfide isomerase family in plants including single-domain protein disulfide isomerase-related proteins. Plant Physiol 137: 762–778
    OpenUrlAbstract/FREE Full Text
  26. ↵
    Iyer VV, Sriram G, Fulton DB, Zhou R, Westgate ME, Shanks JV (2008) Metabolic flux maps comparing the effect of temperature on protein and oil biosynthesis in developing soybean cotyledons. Plant Cell Environ 31: 506–517
    OpenUrlCrossRefPubMed
  27. ↵
    Izui K, Matsumura H, Furumoto T, Kai Y (2004) Phosphoenolpyruvate carboxylase: a new era of structural biology. Annu Rev Plant Biol 55: 69–84
    OpenUrlCrossRefPubMed
  28. ↵
    Kermode AR, Bewley JD (1985) The role of maturation drying in the transition from seed development to germination. J Exp Bot 36: 1906–1915
    OpenUrlAbstract/FREE Full Text
  29. ↵
    Knight JS, Emes MJ, Debnam PM (2001) Isolation and characterisation of a full-length genomic clone encoding a plastidic glucose-6-phosphate dehydrogenase from Nicotiana tabacum. Planta 212: 499–507
    OpenUrlCrossRefPubMed
  30. ↵
    Krepinsky K, Plaumann M, Martin W, Schnarrenberger C (2001) Purification and cloning of chloroplast 6-phosphogluconate dehydrogenase from spinach. Cyanobacterial genes for chloroplast and cytosolic isoenzymes encoded in eukaryotic chromosomes. Eur J Biochem 268: 2678–2686
    OpenUrlPubMed
  31. ↵
    Kruger NJ, von Schaewen A (2003) The oxidative pentose phosphate pathway: structure and organisation. Curr Opin Plant Biol 6: 236–246
    OpenUrlCrossRefPubMed
  32. ↵
    Lara-Núñez A, Rodríguez-Sotres R (2004) Characterization of a dicarboxylate exchange system able to exchange pyrophosphate for L-malate in non-photosynthetic plastids from developing maize embryos. Plant Sci 166: 1335–1343
    OpenUrlCrossRef
  33. ↵
    Miernyk JA, Dennis DT (1992) A developmental analysis of the enolase isozymes from Ricinus communis. Plant Physiol 99: 748–750
    OpenUrlAbstract/FREE Full Text
  34. ↵
    Moorhead GBG, Hodgson RJ, Plaxton WC (1994) Copurification of cytosolic fructose-1,6-bisphosphatase and cytosolic aldolase from endosperm of germinating castor oil seeds. Arch Biochem Biophys 312: 326–335
    OpenUrlCrossRefPubMed
  35. ↵
    Negm FB, Cornel FA, Plaxton WC (1995) Suborganellar localization and molecular characterization of nonproteolytic degraded leukoplast pyruvate kinase from developing castor oil seeds. Plant Physiol 109: 1461–1469
    OpenUrlAbstract
  36. ↵
    Nishimura M, Beevers H (1979) Subcellular distribution of gluconeogenetic enzymes in germinating castor bean endosperm. Plant Physiol 64: 31–37
    OpenUrlAbstract/FREE Full Text
  37. ↵
    Norton G, Harris JF (1975) Compositional changes in developing rape seed (Brassica napus L.). Planta 123: 163–174
    OpenUrlCrossRef
  38. ↵
    Pelzer-Reith B, Penger A, Schnarrenberger C (1993) Plant aldolase: cDNA and deduced amino-acid sequences of the chloroplast and cytosol enzyme from spinach. Plant Mol Biol 21: 331–340
    OpenUrlCrossRefPubMed
  39. ↵
    Petersen J, Brinkmann H, Cerff R (2003) Origin, evolution, and metabolic role of a novel glycolytic GAPDH enzyme recruited by land plant plastids. J Mol Biol 57: 16–26
    OpenUrl
  40. ↵
    Plaxton WC (1991) Leucoplast pyruvate kinase from developing castor oil seeds. Characterization of the enzyme's degradation by a cysteine endopeptidase. Plant Physiol 97: 1334–1338
    OpenUrlAbstract/FREE Full Text
  41. ↵
    Plaxton WC, Podestá FE (2006) The functional organization and control of plant respiration. Crit Rev Plant Sci 25: 159–198
    OpenUrlCrossRef
  42. ↵
    Pleite R, Pike MJ, Garcés R, Martínez-Force E, Rawsthorne S (2005) The sources of carbon and reducing power for fatty acid synthesis in the heterotrophic plastids of developing sunflower (Helianthus annuus L.) embryos. J Exp Bot 56: 1297–1303
    OpenUrlAbstract/FREE Full Text
  43. ↵
    Podesta FE, Plaxton WC (1991) Kinetic and regulatory properties of cytosolic pyruvate kinase from germinating castor oil seeds. Biochem J 279: 495–501
    OpenUrlAbstract/FREE Full Text
  44. ↵
    Rius SP, Casati P, Iglesias AA, Gomez-Casati DF (2008) Characterization of Arabidopsis lines deficient in GAPC-1, a cytosolic NAD-dependent glyceraldehyde-3-phosphate dehydrogenase. Plant Physiol 148: 1655–1667
    OpenUrlAbstract/FREE Full Text
  45. ↵
    Rodríguez-Sotres R, Lara-Núñez A, Rodríguez-Penagos M (2000) Possible roles of phosphoenolpyruvate carboxylase and pyruvate kinase in assimilate partitioning in developing maize embryos. In M Black, K Bradford, J Vazquez-Ramos, eds, Seed Biology: Advances and Applications. CAB International 2000, New York, pp 141–145
  46. ↵
    Ruuska SA, Girke T, Benning C, Ohlrogge JB (2002) Contrapuntal networks of gene expression during Arabidopsis seed filling. Plant Cell 14: 1191–1206
    OpenUrlAbstract/FREE Full Text
  47. ↵
    Sangwan RS, Singh N, Plaxton WC (1992) Phosphenolpyruvate carboxylase activity and concentration in the endosperm of developing and germination castor oil seeds. Plant Physiol 99: 445–449
    OpenUrlAbstract/FREE Full Text
  48. ↵
    Schnarrenberger C, Krüger I (1986) Distinction between cytosol and chloroplast fructose-bisphosphate aldolases from pea, wheat, and corn leaves. Plant Physiol 80: 301–304
    OpenUrlAbstract/FREE Full Text
  49. ↵
    Scholz V, da Silva JN (2008) Prospects and risks of the use of castor oil as a fuel. Biomass Bioenergy 32: 95–100
    OpenUrlCrossRef
  50. ↵
    Schwender J, Ohlrogge JB, Schachar-Hill Y (2003) A flux model of glycolysis and the oxidative pentose phosphate pathway in developing Brassica napus embryos. J Biol Chem 278: 29442–29453
    OpenUrlAbstract/FREE Full Text
  51. ↵
    Shearer HL, Turpin DH, Dennis DT (2004) Characterization of NADP-dependent malic enzyme from developing castor oil seed endosperm. Arch Biochem Biophys 429: 134–144
    OpenUrlCrossRefPubMed
  52. ↵
    Simcox PD, Reid EE, Canvin DT, Dennis DT (1977) Enzymes of the glycolytic and pentose phosphate pathways in proplastids from the developing endosperm of Ricinus communis L. Plant Physiol 59: 1128–1132
    OpenUrlAbstract/FREE Full Text
  53. ↵
    Smith RG, Gauthier DA, Dennis DT, Turpin DH (1992) Malate- and pyruvate-dependent fatty acid synthesis in leucoplasts from developing castor endosperm. Plant Physiol 98: 1233–1238
    OpenUrlAbstract/FREE Full Text
  54. ↵
    Tang GQ, Hardin SC, Dewey R, Huber SC (2003) A novel C-terminal proteolytic processing of cytosolic pyruvate kinase, its phosphorylation and degradation by the proteasome in developing soybean seeds. Plant J 34: 77–93
    OpenUrlCrossRefPubMed
  55. ↵
    Tonge R, Shaw J, Middleton B, Rowlison R, Rayner S, Young J, Pognan F, Hawkins E, Currie I, Davison M (2001) Validation and development of fluorescence two-dimensional differential gel electrophoresis proteomics technology. Proteomics 1: 377–396
    OpenUrlCrossRefPubMed
  56. ↵
    Turner WL, Knowles VL, Plaxton WC (2005) Cytosolic pyruvate kinase: subunit composition, activity, and amount in developing castor and soybean seeds, and biochemical characterization of the purified castor seed enzyme. Planta 222: 1051–1062
    OpenUrlCrossRefPubMed
  57. ↵
    Uhrig RG, O'Leary B, Spang HE, MacDonald JA, She YM, Plaxton WC (2008) Coimmunopurification of phosphorylated bacterial- and plant-type phosphoenolpyruvate carboxylases with the plastidial pyruvate dehydrogenase complex from developing castor oil seeds. Plant Physiol 146: 1346–1357
    OpenUrlAbstract/FREE Full Text
  58. ↵
    Ünlü M, Morgan ME, Minden JS (1997) Difference gel electrophoresis: a single gel method for detecting changes in protein extracts. Electrophoresis 18: 2071–2077
    OpenUrlCrossRefPubMed
  59. ↵
    Valverde F, Ortega JM, Losada M, Serrano A (2005) Sugar-mediated transcriptional regulation of the Gap gene system and concerted photosystem II functional modulation in the microalga Scenedesmus vacuolatus. Planta 221: 937–952
    OpenUrlCrossRefPubMed
  60. ↵
    von Schaewen A, Langenkamper G, Graeve K, Wenderoth I, Scheibe R (1995) Molecular characterisation of the plastidic glucose-6-phosphate dehydrogenase from potato in comparison to its cytosolic counterpart. Plant Physiol 109: 1327–1335
    OpenUrlAbstract
  61. ↵
    Weiss EA (2000) Oilseed Crops, Ed 2. Blackwell Science, Malden, MA, pp 13–51
  62. ↵
    Wendt UA, Wenderoth I, Tegler A, von Schaewen A (2000) Molecular characterisation of a novel glucose-6-phosphate dehydrogenase from potato (Solanum tuberosum L.). Plant J 23: 723–733
    OpenUrlCrossRefPubMed
  63. ↵
    Yates JR, Eng JK, McCormack AL, Schieltz D (1995) Method to correlate tandem mass spectra of modified peptides to amino acid sequences in the protein database. Anal Chem 67: 1426–1436
    OpenUrlCrossRefPubMed
Back to top

Table of Contents

Print
Download PDF
Email Article
Citation Tools
Request Permissions
Quantitative Proteomics of Seed Filling in Castor: Comparison with Soybean and Rapeseed Reveals Differences between Photosynthetic and Nonphotosynthetic Seed Metabolism
Norma L. Houston, Martin Hajduch, Jay J. Thelen
Plant Physiology Oct 2009, 151 (2) 857-868; DOI: 10.1104/pp.109.141622
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

In this issue

View this article with LENS