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BIOCHEMICAL PROCESSES AND MACROMOLECULAR STRUCTURES

Light Enables a Very High Efficiency of Carbon Storage in Developing Embryos of Rapeseed¹

Michigan State University, Department of Plant Biology, East Lansing, Michigan 48824

	ABSTRACT

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
The conversion of photosynthate to seed storage reserves is crucial to plant fitness and agricultural production, yet quantitative information about the efficiency of this process is lacking. To measure metabolic efficiency in developing seeds, rapeseed (Brassica napus) embryos were cultured in media in which all carbon sources were [U-¹⁴C]-labeled and their conversion into CO₂, oil, protein, and other biomass was determined. The conversion efficiency of the supplied carbon into seed storage reserves was very high. When provided with 0, 50, or 150 µmol m^–2 s^–1 light, the proportion of carbon taken up by embryos that was recovered in biomass was 60% to 64%, 77% to 86%, and 85% to 95%, respectively. Light not only improved the efficiency of carbon storage, but also increased the growth rate, the proportion of ¹⁴C recovered in oil relative to protein, and the fixation of external ¹⁴CO₂ into biomass. Embryos grown at 50 µmol m^–2 s^–1 in the presence of 5 µM 1,1-dimethyl-3-(3,4-dichlorophenyl) urea (an inhibitor of photosystem II) were reduced in total biomass and oil synthesis by 3.2-fold and 2.8-fold, respectively, to the levels observed in the dark. To explore if the reduced growth and carbon conversion efficiency in dark were related to oxygen supplied by photosystem II, embryos and siliques were cultured with increased oxygen. The carbon conversion efficiency of embryos remained unchanged when oxygen levels were increased 3-fold. Increasing the O₂ levels surrounding siliques from 21% to 60% did not increase oil synthesis rates either at 1,000 µmol m^–2 s^–1 or in the dark. We conclude that light increases the growth, efficiency of carbon storage, and oil synthesis in developing rapeseed embryos primarily by providing reductant and/or ATP.

Seeds are not simply passive receptacles for the assimilates and minerals provided by the mother plant. They synthesize complex molecules from simple raw materials in relatively precise amounts and proportions (Egli, 1998). Green seeds are photosynthetically active and are able to fix carbon (Watson and Duffus, 1991; Eastmond et al., 1996). As a consequence, the light reactions of photosynthesis potentially can influence the seed's carbon economy, how carbon is stored, and also may play an important role in defining plant fitness and productivity.

In oilseeds, the conversion of carbon supplies into oil may lead to a substantial loss of carbon as CO₂. The primary substrate for oil synthesis in the seeds is the Suc imported from the photosynthetic tissues that is metabolized to generate acetyl-CoA units, the basic building blocks of fatty acids. Acetyl-CoA is formed from pyruvate by an oxidative decarboxylation catalyzed by the pyruvate dehydrogenase complex. In this reaction, for each acetyl-CoA unit produced, one carbon dioxide is also generated, and as a result, 33% of the carbon supplied as to fatty acid synthesis is potentially released as CO₂. This will reduce carbon storage efficiency if mechanisms of CO₂ conservation or recapture in the seeds are lacking. Furthermore, fatty acid synthesis, which primarily occurs in the plastids, requires stoichiometric amounts of ATP, NADPH, and NADH for each sequential addition of an acetyl unit to the growing chain of the fatty acid. In heterotrophic tissues, the plastids must import these cofactors or generate them through carbohydrate oxidation or metabolite shuttles (Rawsthorne, 2002). Several studies with developing rapeseed (Brassica napus) embryos showed that lipid accumulation is stimulated by light (Fuhrmann et al., 1994; Aach and Heise, 1998; Bao et al., 1998; Ruuska et al., 2004), suggesting that fatty acid synthesis may be dependent on the supply of reducing power and ATP provided by the light reactions of photosynthesis. Moreover, Ruuska et al. (2004) calculated that under ambient light conditions, photosynthetic electron transport can meet the demand for NADPH and ATP for rapeseed fatty acid synthesis. Schwender et al. (2004) recently demonstrated that in rapeseed embryos under low light conditions, Rubisco participates in an alternative pathway for carbohydrate to oil conversion that yields 20% more acetyl-CoA and produces 40% less CO₂. Together, these reports suggest that different levels of light may improve the efficiency of carbon storage in green oilseeds like rapeseed.

Despite decades of study of the factors that determine seed yield in crops (Egli, 1998) and the recent improved understanding of pathways involved in carbon supply to fatty acid synthesis in oil-storing seeds (Rawsthorne, 2002; Schwender et al., 2004), we could find no previous study that directly addresses the efficiency by which carbon supplies provided by the mother plant are converted into storage reserves. Such information may be useful for several reasons, including the identification of futile cycles or other potentially inefficient processes that could limit seed yield. Indeed, several studies have indicated that much of the ATP synthesized in heterotrophic plant cells may be consumed by futile cycling (Dieuaide-Noubhani et al., 1995; Rontein et al., 2002; Alonso et al., unpublished data), and we have found that sugars taken up by developing rapeseed embryos undergo potentially wasteful cycling between hexose and triose (Schwender et al., 2003).

To provide a direct measurement of the efficiency of seed carbon metabolism, we developed methods that provide accurate assessment of the conversion of carbon supplies into storage materials and CO₂ by developing rapeseed embryos. Contributions of photosynthesis to the carbon economy were determined at different light levels and by inhibiting PSII using 1,1-dimethyl-3-(3,4-dichlorophenyl) urea (DCMU). Measurements of oil synthesis rates by seeds within detached siliques under different light and oxygen levels were conducted as well. The implications for seed metabolism, physiology, and carbon economy are discussed.

	RESULTS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
Carbon mass balancing is widely used for estimating the efficiency of microorganism performance in processes involving the conversion of substrates into products (Novák and Loubiere, 2000; Converti and Perego, 2002; Sáez et al., 2002). This approach requires that the carbon content of all carbon-containing substrates and products, namely, medium components, biomass, and CO₂ are accurately assessed. To provide a similar assessment of seed carbon use efficiency we took advantage of methods that allow culture of developing embryos in defined media (Smith, 2000). When embryos of rapeseed are dissected from siliques and cultured in medium containing sugars and amino acids (at levels found in endosperm liquid), the embryos develop at rates, and produce storage products, that closely mimic those formed in planta (Schwender and Ohlrogge, 2002). By supplying uniformly ¹⁴C-labeled sugars and amino acids in the medium, and by measuring the ¹⁴C consumption, ¹⁴C incorporation into biomass, and ¹⁴CO₂ production, we could accurately determine carbon balances in developing embryos. In this study, we define the percent carbon conversion efficiency (CCE) as % CCE = ¹⁴C biomass x 100/(¹⁴C biomass + ¹⁴CO₂) (see "Materials and Methods"). We also express carbon use efficiency as the ratio of the moles of carbon incorporated into seed oil to the molar amount of CO₂ developed (oil to CO₂ ratio; ¹⁴C in oil/¹⁴CO₂). Considering that the dominant biosynthetic pathway in rapeseed embryos is fatty acid synthesis and that plastid synthesis of acetyl-CoA from pyruvate is the major source of CO₂ produced, this ratio is a useful measure of the carbon economy of oil synthesis.

Light Increases Embryo Growth and Enables Very High Efficiency of Carbon Metabolism

Table I presents data on ¹⁴C-carbon uptake, biomass accumulation, CO₂ production, oil content, CCE, and the oil to CO₂ ratio after growth of embryos at two light levels and in the dark. Total biomass increase (dry weight) was similar when embryos were grown under 50 or 150 µmol m^–2 s^–1 light. However, in the dark, biomass accumulation was less than one-third that in light. CO₂ production was significantly (P < 0.01) influenced by light levels, decreasing 2-fold between dark and 50 µmol m^–2 s^–1 light and almost 4-fold when light irradiance increased from 50 to 150 µmol m^–2 s^–1. Thus, both increased biomass and reduced CO₂ production suggest that seed metabolism is more efficient under increasing light irradiance. This was further confirmed by the distribution of ¹⁴C-labeling between biomass and CO₂ (Fig. 1). The CCE or proportion of ¹⁴C-labeling in biomass (sum of labeling in oil, protein, and starch + cell wall compounds) increased from 60% in the dark to 86% and 95% of the total ¹⁴C-labeling (¹⁴C biomass + ¹⁴CO₂) at 50 and 150 µmol m^–2 s^–1, respectively. Since the two light treatments (50 and 150 µmol m^–2 s^–1) showed similar ¹⁴C uptake and biomass gain (Table I), the ¹⁴C-label not released as CO₂ is instead incorporated into biomass when light irradiance is increased from 50 to 150 µmol m^–2 s^–1.

View larger version (15K): [in this window] [in a new window]   Figure 1. Distribution of 14C-labeling among biomass fractions and CO2 (100% = 14C biomass + 14CO2): (A) 0 µmol m–2 s–1, (B) 50 µmol m–2 s–1, and (C) 150 µmol m–2 s–1. Embryos were cultured at 21°C for 14 d (at 150 and 50 µmol m–2 s–1) and 21 d (at 0 µmol m–2 s–1) in the presence of U-14C carbon supplies. After culture, the 14CO2 was collected by flushing the flasks through a trapping system, and the embryos were frozen in liquid N2 for analysis of 14C-labeling in biomass components. The percent values shown represent the percentage of 14C recovered in the respective fractions. Because oil, protein, and carbohydrate contain, respectively, 77%, 53%, and 40% carbon by weight, these values differ from weight percent values such as presented in Table I.

Shorter incubations yielded similar results to those observed in the 14- to 21-d experiments presented in Table I and Figure 1. After 3 d of culture, biomass accumulation and lipid synthesis did not differ between the two light levels (50 and 150 µmol m^–2 s^–1) but were 3-fold lower in the dark. CCE was significantly (P < 0.01) increased from 64% in the dark to 77% and 85% at 50 and 150 µmol m^–2 s^–1, respectively. The ratio of carbon stored in oil to carbon released as CO₂ also significantly (P < 0.01) increased with light, from 0.9 to 2.1 to 3.8 at 0, 50, and 150 µmol m^–2 s^–1, respectively. These results confirm that light significantly increases embryo growth and carbon conversion efficiency while reducing loss of carbon as CO₂.

CO₂ Fixation at Different Light Levels

Figure 2 indicates that assimilation of externally supplied ¹⁴CO₂ into biomass increased 3.6-fold when light irradiance was increased from 50 to 150 µmol m^–2 s^–1, suggesting that CO₂ reassimilation by the embryos contributes to the higher carbon conversion efficiency observed at increasing light levels (see above results). Oil labeling at 150 µmol m^–2 s^–1 was 8.4-fold higher than at 50 µmol m^–2 s^–1, whereas protein and cell wall + starch labeling were only 2.8-fold and 2.2-fold higher, respectively, indicating that light irradiance not only increases the total amount of fixed CO₂ but also alters its distribution into different storage products, with an increasing proportion of fixed CO₂ being incorporated into oil at higher light irradiances.

View larger version (14K): [in this window] [in a new window]   Figure 2. Effect of light irradiance on 14CO2 incorporation into biomass. Embryos were grown under a 2% 14CO2 atmosphere (specific activity of C = 44.9 mCi/mol) for 3 d at 150, 50, and 0 µmol m–2 s–1. After culture, the embryos were rapidly frozen in liquid N2, and the 14C-labeling in biomass compounds was measured. A, Total biomass 14C-labeling (microcuries). B, 14C-labeling in oil (microcuries). C, 14C-labeling in protein (microcuries). D, 14C-labeling in starch + cell wall (microcuries). Columns and error bars correspond to the mean ± SD of three independent replicates. Means with the same letter are not significantly different at the P < 0.05 level (Tukey's studentized range test).

Under light conditions, the addition of 5 µM DCMU to the culture medium reduced total biomass accumulation and oil biosynthesis by 3.2-fold and 2.8-fold, respectively, and resulted in DCMU-treated embryos exhibiting values similar to those observed in the dark (Fig. 3). When embryos were cultured in the dark, the presence of DCMU did not significantly affect either total biomass accumulation or oil biosynthesis, indicating that effects of DCMU were specific to light grown cultures (via PSII inhibition) without causing other major effects on embryo metabolism. The CCE and the oil to CO₂ ratio were also reduced by the DCMU treatment in the light, the control (0 µM DCMU) showing 28% and 110% higher CCE and oil to CO₂ ratio, respectively compared to the DCMU treatment. These results indicate that inhibition of PSII significantly reduces biomass and oil synthesis as well as the conversion efficiency of carbon sources into storage products.

View larger version (16K): [in this window] [in a new window]   Figure 3. Effect of inhibition of PSII by DCMU on biomass production, oil synthesis, carbon conversion efficiency, and the oil to CO2 ratio under light and dark conditions. Embryos were cultured in the presence of U-14C carbon sources for 3 d at 50 µmol m–2 s–1 or in the dark with or without DCMU. After culture, the 14C-labeling in the CO2 and biomass compounds was determined. A, Total biomass production expressed in milligrams dry weight (seven embryos). B, Oil synthesis expressed in milligrams (seven embryos). C, CCE, expressed as percent of the sum of 14C biomass and 14CO2. D, Oil to CO2 ratio, calculated from the ratio of 14C-labeling between oil and CO2. Columns and error bars correspond to the mean ± SD of three independent replicates. Means with the same letter are not significantly different at the P < 0.05 level (Tukey's studentized range test).

When oxygen levels in the culture flask were reduced to 1% O₂, embryos in the dark grew little or not at all and CCE was severely reduced to 35% (Fig. 4). Thus, under hypoxic conditions, the efficiency of conversion of assimilates into storage materials is strongly diminished. At superambient O₂ levels (64%), growth in the dark was increased by 30% but CCE did not increase, suggesting that higher oxygen availability in the dark does not increase the efficiency of carbon utilization in the developing embryos.

View larger version (12K): [in this window] [in a new window]   Figure 4. Effect of oxygen on biomass production, oil synthesis, CCE, and oil to CO2 ratio under dark conditions. Embryos were cultured in the dark for 3 d under approximately 1%, 21% (ambient oxygen), and 64% (v/v) O2 in the presence of U-14C carbon sources. A, Total biomass production expressed in milligrams dry weight (seven embryos). B, Oil synthesis, expressed in milligrams (seven embryos). C, CCE expressed as percent of the sum of 14C biomass and 14CO2. D, Oil to CO2 ratio, calculated from the ratio of 14C-labeling between oil and CO2. Columns and error bars correspond to the mean ± SD of three independent replicates. Means with the same letter are not significantly different at the P < 0.05 level (Tukey's studentized range test).

View larger version (15K): [in this window] [in a new window]   Figure 5. Incorporation of radiolabel from (A) 3H2O and (B) [U-14C]Suc into lipids in seeds inside siliques as influenced by light and oxygen levels. A, detached siliques were prelabeled for 24 h (initial) in a buffered solution containing 3H2O. They were then incubated at 1,000 µmol (light treatment) for further 24 h either at ambient air or at 60% oxygen with one-half of the siliques being loosely covered with aluminum foil (dark treatment). After incubation, four seeds from the central region of each silique were removed, the lipids extracted, and the 3H incorporation into fatty acids determined. Columns and error bars correspond to the mean ± SE of 12 independent replicates. Means with the same letter are not significantly different at the P < 0.05 level (Tukey's studentized range test). B, Detached siliques were cultured in the dark in the presence of [U-14C]Suc for 4 d either at ambient air or at 60% oxygen. After incubation, the seeds from each silique were removed, the lipids extracted, and the 14C incorporation into oil determined. Columns and error bars correspond to the mean ± SD of 12 independent replicates. Means with the same letter are not significantly different at the P < 0.05 level (Tukey's studentized range test).

	DISCUSSION

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

Earlier work (Flinn et al., 1977; Pate et al., 1977; Peoples et al., 1985) reported carbon balances conducted in planta in developing fruits of grain legumes that store primarily carbohydrate and protein and little oil. In those studies, the carbon balance was estimated from intake and utilization of carbon and water and from CO₂ exchange measurements. Carbon intake was indirectly deduced from measurements of the amounts of water lost in fruit transpiration during growth, the concentration of solutes delivered to the fruit in xylem and phloem, and the relative ratio of xylem and phloem flows. These previous studies estimated that the conversion of photoassimilates to seed biomass (mainly storage carbohydrates and proteins) is very efficient. In pea (Pisum sativum) fruit, for example, from 18 to 36 d after flowering, almost 80% of the translocate from the parent plant and pod walls was incorporated into dry matter in the seeds (Flinn et al., 1977). Peoples et al. (1985) estimated that 70% of carbon entering the fruit was incorporated into seeds of cowpea (Vigna unguiculata).

Uncertainties about xylem and phloem flow rates and recirculation between the two in those studies (Köckenberger et al., 1997) make these earlier estimates uncertain. Furthermore, these studies and others (e.g. Furbank et al., 2004) have shown that for species such as chickpea, refixation by the fruit wall of CO₂ respired by seeds makes a significant contribution to final carbon recoveries in seeds. For plants with both green seeds and green pod walls, it is difficult to distinguish between contributions of fruit walls and seeds to final seed mass (Willms et al., 1999).

Direct measurement of C uptake and utilization by developing seeds should therefore provide more accurate data for estimating the efficiency of carbon utilization and the factors that influence it. In this study we report a direct analysis of the efficiency of carbon metabolism for developing embryos. Our results indicate that embryo carbon metabolism can be extremely efficient when light is provided, converting up to 95% of carbon input into storage material. For comparison, microorganisms growing aerobically convert approximately 50% of input carbon into biomass (Neidhardt et al., 1990).

Light Increases Biomass Accumulation, Lipid Synthesis, and the Efficiency of Carbon Storage in Rapeseed Embryos

Our results (Figs. 1–3) quantitatively demonstrate the importance of light to achieve high carbon use efficiency in embryos and extend results from several earlier studies. Light may affect carbon utilization by the developing embryos by driving the production of reductant and ATP for fatty acid synthesis. Browse and Slack (1985) concluded that light could provide cofactors for fatty acid synthesis in linseed plastids, and in canola embryos, light stimulates oil synthesis (Fuhrmann et al., 1994; Aach and Heise, 1998; Bao et al., 1998; Ruuska et al., 2004).

The oil to CO₂ ratio is one useful indicator of how efficiently oilseeds use carbon to synthesize oil. As a result of loss of CO₂ in the pyruvate dehydrogenase reaction, conversion of carbohydrate to oil via glycolysis will yield a maximum ratio of two. The values we determined for dark-grown embryos were near one and indicate that the embryos produce CO₂ not only via pyruvate dehydrogenase but also from other respiratory pathways involving carbohydrate oxidation. At 50 µmol m^–2 s^–1, the ratio was higher than two, which can be explained by the involvement of Rubisco and the nonoxidative steps of the pentose phosphate pathway in a metabolic route (the Rubisco bypass) that we recently described (Schwender et al., 2004). The cofactor requirements (ATP and NADPH) of this route are approximately 15% of those required for CO₂ fixation by the Calvin cycle and therefore this route is feasible even with the attenuated light reaching embryos (Schwender et al., 2004). The capacity of the embryos to use light energy even at low irradiances is in agreement with the fact that developing rapeseed seed pigments are adapted to utilize low light in photosynthesis (Ruuska et al., 2004). Plants under field conditions on sunny days receive an average of approximately 1,000 µmol m^–2 s^–1 for up to 12 h. Since the silique wall transmits approximately 20% of incident light, seeds in planta often experience 200 µmol m^–2 s^–1 or more light. At 150 µmol m^–2 s^–1, we observed CCE values approaching 100%, indicating that embryos receiving moderate light are extremely efficient in converting carbon sources into biomass compounds. The very high CCE values observed at 150 µmol m^–2 s^–1 light are higher than predicted by the Rubisco bypass (Schwender et al., 2004). In addition, at 150 µmol m^–2 s^–1, we observed substantial fixation of external CO₂ into oil (Fig. 3), a labeling pattern that is characteristic of the Calvin cycle but not the Rubisco bypass. Together, these observations indicate that cofactor production by the photosystems is sufficient at 150 µmol m^–2 s^–1 to power the Calvin cycle fixation of additional CO₂, thereby increasing the CCE close to 100%.

Increasing light from 50 to 150 µmol m^–2 s^–1 also significantly (P < 0.05) increased the oil to protein ratio. We have observed similar results for Arabidopsis (Arabidopsis thaliana) plants grown under different light levels (Y. Li and J.B. Ohlrogge, unpublished data). These data suggest that in the plastid, greater cofactor production by light can preferentially stimulate fatty acid synthesis or, alternatively, greater phosphoglycerate synthesis via Rubisco may increase substrates availability for plastid fatty acid synthesis. However, light is clearly not essential for high oil production in some seeds such as castor (Ricinus communis) or sunflower (Helianthus annuus) that do not have green seeds. Our initial studies have indicated that isolated sunflower embryos have carbon conversion efficiencies of approximately 60% (data not shown). Other methods of recycling respiratory carbon involving nonseed structures may allow such plants to achieve overall high conversions of photosynthate to seed oil.

Superambient Oxygen Does Not Increase Carbon Storage Efficiency or Lipid Synthesis in Rapeseed

Several recent studies have proposed that seed metabolism may be limited by low oxygen concentrations (Geigenberger, 2003; Vigeolas et al., 2003). In our embryo culture experiments, we found that the rate and composition of biomass accumulation at ambient oxygen are very close to those observed in planta. Vigeolas et al. (2003) reported that developing rapeseed seeds growing in planta at ambient oxygen levels contain approximately 0.8% O₂ (v/v). We attempted to mimic these conditions by growing embryos at low oxygen concentrations. However, the embryos almost stopped growing, accumulating 15-times less biomass than at atmospheric oxygen (Fig. 4A). Thus, an external oxygen level of 1% added to embryo cultures apparently does not reflect in planta conditions, perhaps because the embryos consumed oxygen, resulting in strong hypoxia. Because the silique cavity contains 17% O₂ (Vigeolas et al., 2003), culture of rapeseed embryos under 17% to 21% O₂ likely better represents in planta conditions.

Increasing the oxygen levels from 21% to 64% in the dark did not increase CCE, which remained unchanged at 64%. In addition, for seeds growing inside siliques (Fig. 5), an increase in the oxygen levels from 21% to 60% did not elevate the rate of oil synthesis in the light (1,000 µmol m^–2 s^–1) nor in the dark despite the fact that such an increase in external oxygen levels raises the O₂ levels in the siliques and thus inside the seeds (Vigeolas et al., 2003). Vigeolas et al. (2003) also reported that the lipid synthesis rate in planta increases about 2 times when oxygen levels are increased from atmospheric to 60%. However, the plants in that investigation were cultivated at 300 µmol m^–2 s^–1, whereas in our study the siliques were exposed to 3-fold higher levels of light (1,000 µmol m^–2 s^–1). Seeds and embryos are capable of evolving net oxygen via photosynthesis (Eastmond et al., 1996; King et al., 1998), and under higher levels, light may be sufficient to drive photosynthetic electron transport to provide cofactors and/or oxygen so that oil synthesis is not limited by oxygen availability. The lack of effect of elevated oxygen on embryos inside dark siliques may be because overall metabolism is slower under such conditions and oxygen demand can be met by diffusion from the environment. Whether or not oxygen in fact limits growth and biosynthesis in developing seeds under field light conditions, its utilization for respiration will reduce rather than improve carbon conversion efficiency. Thus, the production of cofactors via photosynthesis is an effective strategy for green seeds to improve carbon storage.

Why is Rapeseed Embryo Metabolism Slow in the Dark?

In this and previous studies, we have observed that rapeseed growth and metabolism are substantially reduced in the dark (Bao et al., 1998; Ruuska et al., 2004; Schwender et al., 2004). Both in culture and in planta embryos have abundant supplies of carbohydrate that could in principle provide cofactors and precursors needed to maintain high rates of metabolism. Thus, it is not immediately clear why metabolism slows at least 2- to 3-fold in the dark. We propose that lowering the rate of storage product accumulation (particularly oil) during the night may be a strategy to reduce carbon lost as CO₂. In rapeseed seeds, an oil content of 45% by weight is equivalent to 61% of seed carbon and therefore over 20% of the imported carbon would be lost at night, whereas in the light, this loss is greatly reduced (Fig. 1).

Based on data of King et al. (1998), the CO₂ produced during the night is unlikely to be recovered by the plant. Even though the sclerenchyma layer of the silique wall may act as a gaseous diffusion barrier causing a build-up of respired CO₂ within the silique locus, King et al. (1998) found that light-dependent CO₂ fixation significantly lowered cavity CO₂ only at 26 DPA, a stage when silique wall chlorophyll is high and seed respiration is low (Eastmond et al., 1996). In addition, at 26 DPA, the CO₂ refixation capacity of the silique wall is 2-fold lower than that of the seed and further decreases to 2.5 times at 30 DPA (King et al., 1998). In developing pea fruits, Flinn et al. (1977) found that more than 40% of ¹⁴CO₂ injected into the fruit gas cavity in the dark was not fixed by pod or seeds, and it was released as ¹⁴CO₂ to the outside atmosphere within 1 h. Together, these studies suggest that in oil plants, the conversion of carbon supplies to reserve materials may cause a large loss of carbon during the night or at very low light irradiances. Thus, green oilseeds like rapeseed may maximize oil synthesis during light periods and strongly reduce oil synthesis under dark conditions to avoid large losses of carbon. This diurnal carbon conservation strategy may ultimately lead to more carbon storage in seeds and thereby, a higher number of successfully established seedlings in the next generation.

	MATERIALS AND METHODS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

Radiochemicals

[U-¹⁴C₆]Glc (317 mCi/mmol, 11.7 GBq/mmol), [U-¹⁴C₁₂]Suc (660 mCi/mmol, 24.4 GBq/mmol), [U-¹⁴C₅]Gln (242 mCi/mmol, 8.95 GBq/mmol), and [U-¹⁴C₃]Ala (162 mCi/mmol, 5.99 GBq/mmol) were from Amersham Biosciences (Piscataway, NJ); NaH¹⁴CO₃ (7.2 mCi/mol, 265.74 GBq/ mmol) was from Sigma-Aldrich (St. Louis).

Plant Material

Rapeseed (Brassica napus) L. cv Reston was grown in 30-cm pots in a greenhouse maintained at 20°C/15°C day/night temperature and with supplemental lighting to provide irradiance of approximately 600 µmol m^–2 s^–1 and a 16-:8-h day/night photoperiod. At 2 weeks, seedlings were thinned to two per pot. Flowers from the main stem were tagged at anthesis and silique development recorded as DPA. Tagged siliques were harvested at 20 DPA and taken to a laminar flow bench for dissection.

Medium Composition and Embryo Culture Conditions

Siliques were surface sterilized with 5% sodium hypochlorite for 10 min and then rinsed three times with sterile water. The developing embryos were dissected under aseptic conditions and transferred into the following culture medium. Carbon and nitrogen medium sources were: Suc (80 mM), Glc (40 mM), Gln (35 mM), and Ala (10 mM). The mineral and vitamin additions were (µg mL^–1): MgSO₄.7H₂O, 370; KCl, 350; CaCl₂.2H₂O, 880; KH₂PO₄, 170; Na₂EDTA, 14.9; FeSO₄.7H₂O, 11.1; H₃BO₃, 12.4; MnSO₄.H₂O, 33.6; ZnSO₄.7H₂O, 21; KI, 1.66; Na₂MoO4.2H₂O, 0.5; CuSO₄. 5H₂O, 0.05; CoCl₂.6H₂O, 0.05; nicotinic acid, 5; pyridoxin hydrochloride, 0.5; thiamine hydrochloride, 0.5; and folic acid, 0.5, respectively. Polyethylene glycol 4,000 was added at 20% to adjust osmotic potential (Schwender and Ohlrogge, 2002), and 50 mM MES was included as a buffer and the medium adjusted to pH 5.8 with 1 N KOH. The embryos were cultured either in a 250-mL screw-cap Erlenmeyer flask sealed with a septum closure or in a regular Erlenmeyer closed with a two-hole rubber stopper connected to two Pasteur pipettes with attached rubber tubing and sleeve stoppers (Fig. 6). The sealing was tested to ensure that the flasks were gas-tight. The septum closure and the sleeve stoppers were included in the culture system to allow CO₂ determinations.

View larger version (22K): [in this window] [in a new window]   Figure 6. Diagram of the culture system used to perform 3-d carbon balances of rapeseed embryos under 2% CO2.

Effect of Light Irradiance on Carbon Balance

Flasks were incubated in a growth chamber at 21°C either under darkness or continuous fluorescent illumination of 50 or 150 µmol m^–2 s^–1. Seven embryos (20 DPA) were cultured in each flask for 14 d. For dark treatment, 12 embryos were cultured for 21 d to compensate for the slow growth in the dark and to provide sufficient biomass for analysis. Uniformly ¹⁴C-labeled carbon supplies (Suc, Glc, Gln, and Ala) were added to the growth medium to provide approximately 10 µCi (370 kBq) total radioactivity per flask. The radioactivity of each compound was adjusted such that each carbon atom added to the growth medium was equally represented at a specific activity of C = 0.89 mCi/mol C. For shorter-term experiments, precultured embryos were transferred to 250-mL Erlenmeyer flasks (7 embryos per flask) and cultured for 3 d in a growth chamber at 21°C either under darkness or light of 50 or 150 µmol m^–2 s^–1 in the presence of radiolabeled carbon supplies as described above.

¹⁴CO₂ Fixation at Different Light Levels

Precultured embryos (7 embryos per flask) were cultured for 3 d under a 2% ¹⁴C-labeled CO₂ atmosphere at three light levels (0, 50, and 150 µmol m^–2 s^–1). The ¹⁴CO₂ was added to headspace by pipetting 100 µL of a NaH¹⁴CO₃ solution in 1 N KOH (100 µCi mL^–1, 3.7 MBq mL^–1) into the internal vial containing unlabel sodium bicarbonate before injecting 1 N HCl. The resulting specific activity of the CO₂ was 44.9 mCi/mol.

Effect of Inhibiting PSII on Carbon Balance

Precultured embryos (7 embryos per flask) were cultured for 3 d under an approximately 2% CO₂ in the presence of radiolabeled carbon sources as described above. The experiment was conducted using a factorial design with light level and DCMU concentration as factors, each with two levels: 0 and 50 µmol m^–2 s^–1, and 0 and 5 µM DCMU, respectively.

Effect of Oxygen on Embryo Carbon Balance

Precultured embryos were incubated in the dark for 3 d under three different O₂ levels (approximately 1%, 21%, and 64% O₂, each with 2% CO₂) in the headspace in the presence of radiolabeled supplies as described above with three replicates per treatment. The gas mixtures were created as follows: approximately 1% O₂ was produced by flushing the flasks with humidified N₂ at approximately 78 mL min^–1 for 9 min 20 s and approximately 64% O₂ by flushing the flasks with humidified O₂ at approximately 78 mL min^–1 for 2 min 4 s. The gases were humidified by bubbling them through water traps placed in series between the gas cylinders and the flasks. Afterward, 6.5 mL of the headspace gas was withdrawn from each flask using a syringe, and the same volume of pure CO₂ was injected into the flasks to create a 2% CO₂ internal atmosphere in all samples. The resulting gas composition of each flask was checked in duplicate analysis at the beginning of the incubation by injecting 100 µL of the headspace gas into the IRGA.

Determination of ¹⁴C-Labeling in Medium, CO₂ Efflux, and ¹⁴CO₂

Immediately after culture, the flasks were placed in an ice bath and 1 mL 0.2 N HCl was injected through the septum or sleeve stopper into the medium to stop metabolism and to release inorganic carbon to the flask headspace. Duplicate 200-µL gas samples were withdrawn from the flask headspace using a 1-mL syringe and injected into an IRGA to determine total CO₂ efflux. The flasks were then flushed for 2 h with nitrogen at 45 mL min^–1 and the exhaust gas bubbled through a 250-mL gas washing bottle containing 140 mL of 1 N KOH. The recovery efficiency of the CO₂ trap was tested with known amounts of NaH¹⁴CO3, being 99.6% (SD% = 3.3). After trapping the CO₂, the embryos were removed and rinsed three times each with 10 mL water to remove surface radioisotope. The embryos were then frozen with liquid nitrogen and lyophilized. The rinse water used for washing the embryos was added to the flasks and the medium was filtered in vacuo through a Buechner funnel with a 55-mm ashless filter paper (fine porosity). The filter and the flask were rinsed several times with water to recover all medium, and the final volume was adjusted to 100 mL. ¹⁴C-label in medium was determined in duplicate analysis by liquid scintillation counting with quenching and background correction. An aliquot (2.5 mL) of the trapping solution containing the ¹⁴CO₂ was pipetted into a 20-mL scintillation vial containing 5 mL of Hionic Fluor scintillation cocktail (Packard BioScience) and counted on a liquid scintillation counter. Analysis was performed in duplicate and all counts were corrected for background and quenching.

Separation of ¹⁴C-Labeled Biomass Compounds

To extract the oil, labeled embryos (approximately 20 mg dry weight) were homogenized with a glass microgrinder at 4°C in 1 mL hexanes:isopropanol (2:1, v/v). The microgrinder tube was centrifuged for 5 min at 5,000g and the supernatant was pipetted into a glass test tube. This extraction was repeated two more times. The pooled lipid fractions were dried under nitrogen at 60°C and redissolved in 4 mL isooctane for scintillation counting. We examined this fraction by thin-layer chromatography (TLC) on silica gel plates developed for 30 min with chloroform/methanol/water (65:25:4 v/v), confirming that it is composed of lipids (>97% of the total counts). After lipid extraction, the pellets were extracted three times each with 1 mL 80% ethanol to recover low-M_r compounds (mostly sugars, amino and organic acids). TLC analysis on cellulose plates of the ethanolic extracts developed with n-butanol/acetic acid/water (4:1:1 v/v) for 2 h (three times) using U-¹⁴C standards (Suc, Glc, Gln, and Ala) confirmed that this fraction mainly contains the carbon sources included in the culture medium (data not shown). The remaining pellet was then dried at room temperature under a stream of nitrogen and resuspended in 1 mL of 0.01 M sodium phosphate saline buffer, pH 7.4, containing 1 mM EDTA, 10 mM 2-mercaptoethanol, 0.02% (w/v) sodium azide,l and 0.0125% (w/v) SDS. The homogenate was transferred to a glass tube and 3 mL buffer solution along with four glass beads (4-mm diameter) were added to the samples. The proteins were extracted by mixing the samples with a Vortex mixer for 15 min. The tubes were then centrifuged at 3,200g for 15 min and the supernatants transferred into another glass tube. The proteins were further extracted by repeating this procedure after adding an additional 4 mL of buffer. The collected protein extracts were pooled and an aliquot sampled for scintillation counting. The pellet was then dried under nitrogen at 60°C and 0.5 mL of 67% (v/v) aqueous sulfuric acid was added to the samples to hydrolyze starch and cell wall components. The pellet was mixed with a Vortex mixer and incubated for approximately 5 min at 60°C until samples turned slightly red. The reaction was stopped by adding 4.5 mL of 1 N KOH and an aliquot of the suspensions was counted. ¹⁴C-label was determined in duplicate analysis in each biomass fraction (oil, protein, and starch and cell wall components) by liquid scintillation counting with quenching and background correction.

Determination of Oil Content

Total oil content of embryos was determined by gas chromatography with flame ionization detection of fatty acid methyl esters. One milliliter of the above lipid extract together with 1 mL of a 750 µg mL^–1 triheptadecanoin solution in isooctane (internal standard) were dried under nitrogen at 60°C, and then transmethylated for 45 min at 60°C with 1 mL of a 0.5-M solution of sodium methylate in methanol. Isooctane (1.5 mL) and 0.5 mL of 5% (w/v) NaHSO₄ in water were added with mixing and tubes centrifuged for 10 min at 3,200g. One microliter of the isooctane phase was analyzed on a DB-23 capillary column (30-m x 0.25-mm i.d. x 0.25-µm thickness; Agilent J&W, Palo Alto, CA). The carrier gas was helium at a pressure of 120 kPa. The oven temperature was programmed as follows: the initial temperature (150°C) was kept for 3 min, increased linearly to 210°C at a 5°C/min rate, and then increased to 250°C at 20°C/min, being the final temperature hold for 3 min. The injector and detector temperatures were 270°C and 280°C, respectively. The samples were injected at a split rate of 15:1.

Calculation of Carbon Balance, Carbon Recovery, and Carbon Conversion Efficiency

The carbon balance of rapeseed embryo cultures was determined by providing all carbon sources uniformly ¹⁴C-labeled at the same specific activity and determining ¹⁴C incorporation into biomass end products, the ¹⁴C radioactivity remaining in the growth medium and released as ¹⁴CO₂. Carbon uptake was estimated as the difference between initial ¹⁴C-labeling (10 µCi, 370 kBq) minus final ¹⁴C-labeling present in the medium after culture. In preliminary experiments, NMR and TLC analysis of the medium after culture of embryos (data not shown) indicated that embryos do not secrete products (e.g. malate) at levels that would alter the carbon balance. The recovery of carbon after culture (carbon recovery [percent] = [¹⁴C biomass + ¹⁴CO₂] x 100/¹⁴C uptake) was 99.5% (SD = 5.3%), which indicates that all the major carbon components in the system are properly accounted for. Therefore, the sum of measurements of ¹⁴C incorporation into embryo biomass plus ¹⁴CO₂ represents the total uptake of labeled carbon supplies. ¹⁴C-labeled biomass was calculated as the sum of each biomass fraction (¹⁴C biomass = ¹⁴C oil + ¹⁴C protein + ¹⁴C starch and cell wall). The 80% ethanolic fraction (<15% of total ¹⁴C in 3-d experiments) was not included in this calculation because it largely represents substrate uptake but not carbon conversion to storage products. CCE was calculated as: percent CCE = ¹⁴C biomass x 100/[¹⁴C biomass + ¹⁴CO₂].

Effect of Oxygen on Oil Synthesis in Seeds within Siliques

Lipid Labeling Using ³H₂O as Precursor
Because it equilibrates rapidly with cellular water and exchanges ³H with hydrogen atom of NAD(P)H that participate in reduction steps of fatty acid synthesis, ³H₂O is an effective tracer for lipid synthesis (Bao et al., 1998; Ruuska et al., 2004). To assess the influence of oxygen on seeds within siliques, ³H₂O was fed to the seeds via the transpiration stream as follows. Eighteen siliques at 30 DPA were removed from greenhouse grown plants and the pedicle was immediately placed into a solution of 5 mCi/mL ³H₂O, 50 mM Suc, 50 mM HEPES, pH 5.7, and 3 mg/mL Murashige and Skoog basal salts (Murashige and Skoog, 1962). After 24 h in the light to allow movement of water into the silique (and its seeds) via transpiration, seeds from six siliques were analyzed for ³H prelabeling into oil as described below. The remaining 12 siliques were further incubated for 24 h in two 1-L jars continuously flushed (1 L min^–1) with either ambient air or with 60% oxygen. Increasing ambient oxygen surrounding siliques has been shown to rapidly increase silique locule oxygen content (Porterfield et al., 1999). Light for each jar was provided from two 500-W projector bulbs shining through CuSO₄ solution in Blake bottles to absorb infrared light. Light impinging on each silique was approximately 1,000 µmol m^–2 s^–1. Temperature in the jars did not increase during the experiment. One-half of the siliques in each jar were loosely covered with aluminum foil with both ends open and allowing air space surrounding the silique. During the entire 48-h experiment, approximately 1 mL of solution was taken up (transpired) by each silique. Four seeds from the central region of each silique were then removed, placed into individual screw cap tubes, and heated with 0.1 mL isopropanol for 10 min at 80°C. After cooling, seeds were homogenized and 0.15 mL hexane added and allowed to extract overnight at room temperature. After adding 0.8 mL hexane and 0.2 mL 0.1 M acetic acid, 0.35 mL of the upper phase lipid extract was transferred to a scintillation vial, evaporated at 70°C, and counted to determine ³H incorporation into fatty acids.

Lipid Labeling in the Dark Using [U-¹⁴C]Suc as Precursor
We performed an additional experiment using [U-¹⁴C]Suc as lipid precursor with no prelabeling. Siliques at 23 DPA were taken from greenhouse plants and surface sterilized as above. The siliques' pedicels were placed into 15-mL vials containing 150 mM [U-¹⁴C]Suc (10 µCi, 370 kBq), 70 mM Gln, and 4.3 mg mL^–1 Murashige and Skoog basal salts, pH 5.8. Three vials with two siliques per vial were placed inside 1-L screw-cap Erlenmeyers covered with aluminum foil. Two flasks per oxygen treatment were sealed after transferring the vials inside the flasks. A 60% O₂ (v/v) atmosphere was created by flushing the flasks with humidified O₂ at approximately 78 mL min^–1 for 10 min 16 s. The gas composition was checked by injecting 100 µL of the flask gas headspace into the IRGA. After 4 d of labeling at 21°C, the siliques were then removed from the vials and immediately submerged into liquid nitrogen and lyophilized for 48 h. The seeds from each silique were removed, placed in 1.5-mL polypropylene vials containing a 6 mm-stainless steel ball, and homogenized in 0.5 mL 2:1 (v/v) hexanes:isopropanol. After centrifugation, lipids were extracted two more times each with 1 mL of solvent mixture. Pooled extracts were directly dried inside scintillation vials under N₂ at 60°C. Five milliliters of scintillation cocktail were added to each vial and ¹⁴C-labeling determined by liquid scintillation counting.

Statistics

One-way ANOVA and Tukey's studentized range test were performed using the general linear model procedure of SAS statistical software (SAS Institute, NC). Unless otherwise indicated, all experiments were done with three replications per treatment.

	ACKNOWLEDGMENTS

 
We thank Prof. Randy Beaudry for helpful advice in the determination of gas composition and Dr. Mike Pollard for kindly providing us his protocol for TLC of lipids.

Received March 31, 2005; returned for revision May 13, 2005; accepted May 16, 2005.

	FOOTNOTES

 
¹ This work was supported by the National Science Foundation (grant no. MCB 0224655) and by the U.S. Department of Agriculture (grant no. 2003–35321–12935). Acknowledgment is also made to the Michigan Agricultural Experiment Station for its support of this research.

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.063628.

^* Corresponding author; e-mail goffman{at}msu.edu; fax 517–353–1926.

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