- © 2016 American Society of Plant Biologists. All rights reserved.
Abstract
C4 photosynthesis in grasses requires the coordinated movement of metabolites through two specialized leaf cell types, mesophyll (M) and bundle sheath (BS), to concentrate CO2 around Rubisco. Despite the importance of transporters in this process, few have been identified or rigorously characterized. In maize (Zea mays), DCT2 has been proposed to function as a plastid-localized malate transporter and is preferentially expressed in BS cells. Here, we characterized the role of DCT2 in maize leaves using Activator-tagged mutant alleles. Our results indicate that DCT2 enables the transport of malate into the BS chloroplast. Isotopic labeling experiments show that the loss of DCT2 results in markedly different metabolic network operation and dramatically reduced biomass production. In the absence of a functioning malate shuttle, dct2 lines survive through the enhanced use of the phosphoenolpyruvate carboxykinase carbon shuttle pathway that in wild-type maize accounts for ∼25% of the photosynthetic activity. The results emphasize the importance of malate transport during C4 photosynthesis, define the role of a primary malate transporter in BS cells, and support a model for carbon exchange between BS and M cells in maize.
INTRODUCTION
C4 photosynthesis comprises a series of metabolic reactions that exploit specialized cell types and biochemistry to concentrate CO2 around Rubisco (Hatch and Slack, 1966; von Caemmerer and Furbank, 2003). Under hot dry conditions, C4 plants maintain high rates of biomass production with improved water and nitrogen use efficiency relative to many C3 species (Leegood et al., 1996; Sage, 2004; Byrt et al., 2011). Apart from a few exceptions (Edwards et al., 2004; King et al., 2012), the photosynthetic activities of C4 plants are coordinated between two specialized leaf cell types, the inner bundle sheath (BS) and the outer mesophyll (M), arranged in concentric rings around the vasculature in a pattern known as Kranz anatomy. C4 plants are traditionally classified into three distinct biochemical subtypes, each named after the enzyme that performs the primary decarboxylation reaction in the BS cell: chloroplastic NADP-dependent malic enzyme (NADP-ME), mitochondrial NAD-dependent malic enzyme (NAD-ME), and cytosolic phosphoenolpyruvate carboxykinase (PEPCK) (Hatch, 1975). Increasingly, the evidence suggests that these pathways operate in concert in many species (Hatch, 1971; Chapman and Hatch, 1981; Furbank, 2011; Pick et al., 2011; Wang et al., 2014b). For example, the NAD-ME subtype Cleome gynandra and the NADP-ME species maize (Zea mays) both contain significant amounts of active PEPCK in BS cells (Walker et al., 1997; Muhaidat et al., 2007; Sommer et al., 2012; Muhaidat and McKown, 2013). When isolated maize BS strands were incubated with aspartate, they generated CO2 (Chapman and Hatch, 1981) and phosphoenolpyruvate (PEP) (Wingler et al., 1999), consistent with the function of a PEPCK pathway that supplies CO2 to the Calvin cycle and regenerates PEP to sustain further rounds of C4 metabolism. Additionally, studies at the gene and metabolite level including RNA and metabolite profiling experiments in maize provide further evidence for the concomitant operation of NADP-ME and PEPCK decarboxylation pathways (Li et al., 2010; Pick et al., 2011; Wang et al., 2014a)
The partitioning of photosynthetic activities between the BS and M cells in C4 plants requires an extensive intercellular and intracellular exchange of metabolites through plasmodesmata and organelle membranes, respectively, regardless of the C4 subtype pathway in use (Bräutigam et al., 2008). Carbon movement is very rapid and significantly exceeds assimilation rates (Laisk and Edwards, 2000), requiring the close coordination of enzyme activities with cell- and organelle-specific export and import. This rapid transport across the mesophyll chloroplast envelope has been validated for all four metabolites (oxaloacetate [OAA], malate, pyruvate, and PEP) involved in the core C4 carbon shuttle pathway (Huber and Edwards, 1977a, 1977b; Hatch et al., 1984; Flugge et al., 1985), and when exchange is impaired the rate of photosynthesis is severely limited (Bräutigam and Weber, 2011). The first step in the C4 cycle is the assimilation of one molecule of CO2 into OAA by PEP-carboxylase, which is subsequently transported into the M chloroplast and reduced to malate by a plastid-localized NADP-dependent malate dehydrogenase. Malate is exported from the chloroplast and diffuses into the BS cytoplasm (Stitt and Heldt, 1985; Wang et al., 2014a) prior to import into the BS chloroplast. The BS chloroplast contains Rubisco and several Calvin-Benson cycle enzymes that assimilate CO2 produced from malate decarboxylation by NADP-ME. The other product of decarboxylation, pyruvate, is used to regenerate PEP that is needed for repeated cycles of C4 photosynthesis. The movement of pyruvate and PEP requires at least two transport steps to import and export three carbon molecules between the BS and M plastids to complete the NADP-ME cycle. Transporters are also necessary to export 3-phosphoglyceric acid (3-PGA) for reduction to glyceraldehyde 3-phosphate (or triose phosphate) in the M cells, which is then returned to the BS chloroplast (Weber and von Caemmerer, 2010). In total, at least 10 import and export steps are needed per CO2 molecule assimilated. Although extensive use of transporters is crucial for the proper function of C4 photosynthesis, elucidation of the specific proteins has proven challenging (Bräutigam et al., 2008; Majeran and van Wijk, 2009; Weber and von Caemmerer, 2010).
Three dicarboxylate transporters that can efficiently move malate have been identified in maize. DICARBOXYLIC ACID TRANSPORTER1 (DCT1), DCT2, and 2-OXOGLUTARATE/MALATE TRANSPORTER1 (OMT1) (Taniguchi et al., 2004) are predicted to be chloroplast localized (Taniguchi et al., 2004; Majeran and van Wijk, 2009) and differentially expressed between the M and BS cells (Li et al., 2010; Chang et al., 2012; Tausta et al., 2014), but their metabolic function to date is unclear. The DCT2 protein and transcript are enriched in the BS of maize (Bräutigam et al., 2008; Li et al., 2010), and expression profiles across the developmental leaf gradient coincide with other photosynthetic genes (Li et al., 2010). DCT2 transcript levels also respond to light exposure (Taniguchi et al., 2004; Li et al., 2010), which supports their involvement in photosynthesis.
To functionally assess the role of DCT2 in maize, we used Activator (Ac) insertional mutagenesis to create mutant alleles of DCT2. The homozygous loss-of-function alleles displayed a pale-green/yellow mutant phenotype and resulted in reduced viability. Under low-light conditions in a greenhouse, mutants could be nurtured to maturity, occasionally resulting in a limited number of seeds from homozygous plants. Transport assays, transcript profiling, transmission electron microscopy, and multiple 14C, 13C, and 2H isotopic labeling experiments described in this article indicate that the reduced growth is likely a consequence of impaired malate transport into the BS chloroplast of dct2 lines. Labeling experiments further support the operation of a functional PEPCK type pathway in maize that can sustain limited growth when malate transport is compromised. The results provide biochemical evidence for the operation of two C4 subtype pathways concomitantly and describe the role of a key transport step in C4 photosynthesis.
RESULTS
Generation of DCT2 Mutant Lines by Targeted Ac Mutagenesis
Mutant alleles of DCT2 were recovered through insertional mutagenesis that used the Ac transposable element (McClintock, 1948). Transposition events containing an insertion into or around DCT2 were identified by reverse genetics using bti99221::Ac as a donor transposon that was close in physical and genetic proximity to DCT2 (401.2 kb, 0.44 cM). Selections were performed for increased copy number of the Ac element that enrich for transposition events as previously described (Brutnell and Conrad, 2003; Kolkman et al., 2005). Genetic screens of 1050 progeny identified four insertion sites that were fine mapped to the first exon (dct2-1::Ac and dct2-2::Ac), the 5′ untranslated region (dct2-3::Ac), and the second intron (dct2-4::Ac) of DCT2 (Figure 1A). Self-pollination resulted in homozygous lines for further analysis (referred to hereafter as dct2 without the Ac designation). Plants, homozygous for insertions dct2-3 and dct2-4, did not show a mutant phenotype and were deemed too far upstream to affect DCT2 transcription (dct2-3) or were efficiently spliced out of the transcript (dct2-4). Nine-day-old plants, homozygous for the dct2-1 and dct2-2 alleles, were phenotypically similar, with pale-yellow leaves (Figures 1B and 1C), low net photosynthetic rates (Figure 1F), and low chlorophyll content (Figure 1G). The mutants lacked the typical yellow-to-green color gradient observed in developing wild-type leaves (Figure 1C) and had low concentrations of Calvin cycle intermediates (Supplemental Table 1). In addition, quantitative real-time PCR failed to detect DCT2 transcripts in homozygous dct2-1 and dct2-2 lines (Figure 1E). Plants homozygous for these strong loss-of-function alleles developed slowly, resulting in thinner leaves and shorter plants that produced less biomass and infrequently generated seeds (Figure 1D). The metabolic features of dct2-1 were further characterized to assess the molecular basis underlying these phenotypes.
Molecular and Physiological Data for Wild-Type and dct2 Lines.
Insertional mutagenesis of DCT2 resulted in pale leaves and loss of carbon assimilation gradient.
(A) Ac.bti99221, located 401.2 kb (0.44 cM) 5′ of DCT2, relocated creating four insertions in and around the gene: two in the first exon (dct2-1 and dct2-2), one in the 5′ untranslated region (dct2-3), and one in the second intron (dct2-4). Red triangles indicate the 5′ end of the Ac insert.
(B) Wild-type (WT) and mutant plants (dct2-1 and dct2-2) 9 d after sowing. Red demarcations indicate a metric ruler with labeled centimeter increments.
(C), (F), and (G) Nine-day-old leaves of the wild type and dct2-1 showing loss of color gradient. Sections A, B, and C were tested for net CO2 assimilation (F) with an infrared gas analyzer (LI-6400). Segments −4, −1, +4, and +10 were sampled for RNA-seq and metabolite quantification, including chlorophyll levels (G).
(D) Five-week-old wild-type and dct2-1 and dct2-2 plants showing the impeded development of mutant plants compared with the wild-type.
(E) Quantitative real-time PCR of DCT2 transcripts in the wild type (black). dct2-1 (white) and dct2-2 (gray) show no detectable transcripts of DCT2. Target/ref ratio calculated as DCT2/tubulin expression level. Error bars in (C), (E), and (F) represent sd (n = 3).
Leaf Development Is Impaired in Homozygous dct2-1 Plants
To characterize the phenotype of the mutant plants at the cellular level, histological surveys of dct2-1 and wild-type plants were assessed by transmission electron microscopy (TEM). Samples from four segments were collected along a developing third leaf at 9 d after sowing, as previously described (Li et al., 2010). The segments were referenced relative to the position of the ligule of leaf 2, as this point represents the source-sink boundary in a wild-type leaf, and demarcates the transition from shaded to fully exposed leaf tissue. Thus, segment −4 was defined by location at the base of the leaf blade that is photosynthetically inactive, −1 was taken at the sink-to-source transition zone (just below the point of leaf emergence from the whorl), segment +4 was from the mid-leaf, a region that is exposed to light and photosynthetically active (maturation zone), and +10 was from the tip of the leaf that is the terminally differentiated tissue (Figure 1C). The ultrastructure of the dct2-1 leaf in all segments suggested delayed development compared with wild-type plants (Figure 2). In the −4 segment of the leaf, both BS and M cells of dct2-1 had a large number of vacuoles (Figures 2C and 2D), indicating a lag in development compared with the fused vacuoles in wild-type plants (Figures 2A and 2B). In addition, dct2-1 BS plastids were small and regularly dividing, which is consistent with hindered leaf development and a sign of an immature tissue (Figures 2C and 2D). In the −1 segment, the chloroplasts in both BS and M cells of dct2-1 were also smaller with poorly developed granum lamellae (Figures 2E to 2H). In the +4 segment, both the wild type and dct2-1 showed the typical NADP-ME centrifugal arrangement of plastids. The thylakoid membrane of wild-type chloroplasts thickened in both BS and M cells, and the BS chloroplasts exhibited a small degree of granal stacking (Figures 2I and 2J). dct2-1 BS chloroplasts remained round, similar to their shape at the −1 segment, and in both BS and M chloroplasts, the thylakoid membranes did not show a significant thickening compared with the −1 segment (Figures 2K and 2L). Chloroplasts in both BS and M cells from the wild type were large by the +10 segment and showed an accumulation of starch granules and stacking in BS chloroplasts (Figures 2M and 2N), whereas those from dct2-1 remained in the same state as in the +4 segment and did not have starch granules or display granal stacks (Figures 2O and 2P). This impaired plastid development in dct2-1 plants may be a consequence of carbon starvation or due to a defect in related metabolic pathways.
TEM of Wild-Type and dct2-1 Leaf Segments.
TEM analysis of the ultrastructure of four segments sampled along a developing third leaf at 9 d after sowing suggests impeded development of photosynthetic cells in dct2-1. The four segments correspond to four rows of panels in the figure and indicate position along the leaf inspected from the base of the leaf (also called −4) to the tip (also called +10) as stated along the y axis. For both the wild type and dct2-1, two columns are presented that show varying degrees of magnification to emphasize salient features described in the text. In particular, the panels emphasize a differing number of vacuoles in −4, differing grana structure and chloroplast size in −1 and +4, and reduced starch and granal stacking in +10. Ch, chloroplast; Vc, vacuole; sg, starch. Bars = 20 µm in (A), (G), (I), (K), (M), and (O), 1 µm in (B), (H), and (J), 10 µm in (C) and (E), and 2 µm in (D), (F), (L), (N), and (P).
DCT2 Transports Malate into the BS Chloroplast during C4 Photosynthesis
DCT2 encodes a passive-mediated transporter that is minimally capable of exchanging malate for 2-oxoglutarate (2-OG), glutamate, or oxaloacetate (Taniguchi et al., 2004). The malate transport activities of BS chloroplasts were compared between wild-type and dct2-1 plants (Figures 3A and 3B). Due to low-yield and low density of intact dct2-1 BS chloroplasts, the direct uptake of malate into this organelle could not be assessed with traditional silicone-oil centrifugation fractionation methods. However malate-dependent pyruvate formation by maize BS chloroplasts has previously been described (Boag and Jenkins, 1985), as an indirect assay of malate import activity by measuring the conversion of exogenous malate to pyruvate in BS chloroplasts with a stromal NADP-ME. Organelle preparations using a Percoll gradient resulted in ∼89% wild-type and 83% dct2-1 intact isolated chloroplasts, respectively, as indicated by NADP-ME activity before and after chloroplast lysis (Figure 3B). Chlorophyll content per chloroplast was approximately 3 times lower in dct2-1 than in wild-type plastids. Therefore, activities were normalized by chloroplast number (µmol [109 chloroplasts]−1 h−1), and then the transport capacity was compared relative to maximal NADP-ME activity from intact chloroplasts (Figure 3B). This assay revealed that pyruvate was produced at a slower rate in dct2-1 chloroplasts relative to the wild type. Furthermore, chloroplast lysis with detergent resulted in elevated pyruvate formation rates for both the wild type and dct2-1, suggesting a significant transport limitation in both lines (Boag and Jenkins, 1985). Comparison of the lysed chloroplast values using either the NADP-ME activity assay (i.e., 846/1828; ∼46%) or pyruvate formation (i.e., 391/826; ∼47%) measurements showed a similar decrease in activity in dct2-1. The normalized pyruvate formation rate was a modest 2.3% (wild type) or 0.7% (dct2-1) relative to NADP-ME activity with the wild type value being comparable to other reports (Boag and Jenkins, 1985; Jenkins and Boag, 1985; Boag and Jenkins, 1986). The limited pyruvate formation observed in dct2-1 plants was likely due to either some malate import or leakage of the isolated plastids. Regardless, the assay demonstrated compromised malate import associated with the dct2-1 mutation. Malate-dependent pyruvate formation generated from wild-type chloroplasts was enhanced in the presence of aspartate (Figures 3A and 3B), as has been previously reported (Boag and Jenkins, 1985, 1986; Jenkins and Boag, 1985), but the enhancement was not observed in the mutant chloroplasts, suggesting that aspartate is unable to stimulate a malate-specific import process in dct2 plants (discussed further below).
Malate Transport in the Wild Type and dct2-1.
Malate-dependent pyruvate formation, NADP-ME activities, and 2,3,3-2H-malate labeling of BS chloroplasts were measured in 9-d-old wild-type and dct2-1 plants.
(A) Time course of malate-dependent pyruvate formation in BS chloroplasts of wild-type (square symbols) and dct2-1 (round and triangle symbols) plants. BS chloroplasts were suspended in 10 mM malate with or without 10 mM aspartate and the amount of pyruvate formed was quantified spectrophotometrically (se, n = 3).
(B) NADP-ME activity and pyruvate (PYR) formation rate in BS chloroplasts of wild-type and dct2-1 plants (se, n = 3). NADP-ME activity was based on spectral detection of NADP at 340 nm. The rates of pyruvate formation in chloroplasts with or without aspartate were calculated from the slopes in (A) (2 to 5 min). Calculations for the intact chloroplast NADP-ME activity, chloroplast intactness (%), and pyruvate formation in intact chloroplasts normalized to NADP-ME activity (%) are defined in the gray box. Intact chloroplast NADP-ME activity indicates the formation of pyruvate that is not transport limited; thus, normalization by this value provides a comparison specific to the transport function. The percentages in parentheses for pyruvate formation from lysed chloroplasts indicate the pyruvate formation rate relative to maximal NADP-ME activity (e.g., 826/1828; 45%).
(C) Quantification of malate labeling in BS chloroplasts after 1 h of incubation with 2,3,3-2H-malate (purple box, deuteriums indicated by “D” in the small red boxes). LC-MS/MS chromatograms indicate the peaks of unlabeled ([M]+) and triple labeled ([M+3]+) malate. Malate pool sizes in dct2-1 BS chloroplasts was 42% of the wild type; therefore, twice the amount was injected onto LC-MS/MS for labeling quantification in the spectra. Table: fractional abundance of mass isotopomers in 2,3,3-2H-malate standard, wild-type, and dct2-1 BS chloroplasts after labeling (sd, n = 3). [M+i]+, the ith-labeled mass isotopomer. Labeling data were corrected for natural isotope abundance.
Malate uptake into the BS chloroplasts was further characterized with 2,3,3-2H malate that was supplied to isolated wild-type and dct2-1 BS chloroplasts (Figure 3C). Because the stable isotope incorporation is relative to the entire unlabeled pool, mass spectrometry can judge the absolute amount of malate transport relative to its pool size and additional normalizations were unnecessary. The measurement of 2H-malate was therefore a dynamic, complementary evaluation of metabolism relative to enzyme assays, though requiring additional isolated dct2-1 chloroplasts for sensitive measurement. Significantly more triply labeled malate (malate with three hydrogens replaced by three deuteriums [M+3]+) was present in chloroplasts from wild-type (60.3% ± 0.8%) than dct2-1 plants (18.7% ± 4.0%) (Figure 3C) and was qualitatively comparable to the NADP-ME activity assays. The levels of malate in BS cells and in isolated chloroplasts were measured to check that the differences in labeling were not merely a consequence of the preexisting pools. The level of malate in washed chloroplasts was ∼1% of the level in BS cells for both the wild type and dct2-1, consistent with other subcellular measurements of malate (Weiner and Heldt, 1992); however, dct2-1 chloroplasts had reduced malate concentrations (0.49 ± 0.01 nmol gFW−1 BS) relative to the wild type (1.16 ± 0.20 nmol gFW−1 BS). The smaller pools in dct2-1 would be expected to turnover more quickly with isotope (i.e., they would be more labeled) if the transport rate indeed was similar to that of the wild type. Since this was not observed, the differences in isotope incorporation were a consequence of impaired malate transport in dct2-1 chloroplasts and not of changes in pool size. Based on the calculated absolute incorporation of malate labeling in the pool of the wild type and dct2-1, direct malate movement into the chloroplast in dct2-1 could account for ∼13% of the wild-type activity, but could also reflect leakiness in the isolated chloroplast system.
dct2-1 Plants Have Reduced Calvin Cycle Flux Relative to the Wild Type
Metabolic changes in the Calvin-Benson cycle and related pathways that resulted from the loss of DCT2 function were characterized with carbon isotope labeling experiments. Air containing 13CO2 substituted for 12CO2 at near ambient levels (∼330 ppm) was provided to leaf segments (i.e., proximal one-third of the leaf from the tip; sections B+C in Figure 1C) of 9-d-old wild-type and dct2-1 plants, resulting in the labeling of primary intermediates including sugar phosphates, amino acids, and organic acids. Mass isotopomers were quantified with liquid chromatography-tandem mass spectrometry (LC-MS/MS) and gas chromatography-mass spectrometry (GC-MS) of samples extracted at 7 to 11 time points during the labeling period. The average 13C enrichment of each metabolite was calculated after correction for natural abundance. 13C labeling was more pronounced in Calvin cycle intermediates, including 3-PGA, ribulose-1,5-bisphosphate, and sedoheptulose-7-phosphate from the wild type relative to dct2-1 (Figure 4) and approached isotopic steady state within 3 min (∼85% 13C). Similar levels of isotope incorporation have been observed in C3 plants (Ma et al., 2014), though requiring a longer labeling duration (∼15 min). By contrast, 13C enrichment was limited (∼20% 13C) in the dct2-1 line. Metabolites that are derived from 3-PGA, including pyruvate, PEP, and alanine (Supplemental Figure 1), were also labeled more rapidly in the wild type and reflected the asymmetrical isotopic pattern of Rubisco-based assimilation (Supplemental Figure 1B). In the wild type, the combined enrichment of all three carbons from alanine was greater than a measurement of the second and third carbons, indicating that the first carbon was most highly labeled through the Calvin cycle. This labeling description is consistent with 13C incorporation into the first carbon position of 3-PGA through photosynthetic assimilation.
13CO2 Labeling of Metabolites.
Data were collected from 13CO2 (∼330 ppm) labeled leaf segments (sections B+C in Figure 1C) of 9-d-old wild-type (solid symbols) and dct2-1 plants (open symbols) at a light intensity of 500 µmol m−2 s−1 and analyzed for labeling of Calvin cycle intermediates including 3-PGA (A), ribulose-1,5-bisphosphate (RuBP) (B), and sedoheptulose-7-phosphate (S7P) (C). Labeled samples were collected over a 3-min interval at 20, 40, 60, 90, 120, 150, and 180 s. Average 13C enrichment (%) was calculated using the formula defined in the gray box, where N is the number of carbon atoms in the metabolite and Mi is the fractional abundance (%) of the ith mass isotopomer. Mi data were corrected for natural isotope abundance (sd, n = 3). Additional mass isotopomer data are reported in Supplemental Data Set 2.
Malate and Aspartate Labeling Indicate Altered C4 Subtype Pathway Activities
The simultaneous use of malate and aspartate as substrates for carbon concentration in the BS occurs in both NADP-ME and PEPCK C4 plants (Hatch, 1971; Chapman and Hatch, 1981; Furbank, 2011; Pick et al., 2011; Wang et al., 2014a). Modeling efforts have also suggested coordination between coexisting C4 pathways (Wang et al., 2014b). Here, the movement of carbon in wild-type maize and dct2-1 plants was tracked by quantifying the absolute 13C incorporation into malate and aspartate pools over the 3-min 13CO2 time course (Figure 5). 13CO2 incorporated with PEP by PEP-carboxylase resulted in singly labeled OAA that is converted to either malate or aspartate (Figure 5A). The relative abundance (%) of the singly labeled mass isotopomer [M+1]+ (M, the base mass; 1, one carbon labeled) in malate and aspartate (Figure 5B) was combined with pool size information (Figure 5C) to determine the initial rate of labeling of these metabolites (Figure 5D). The slopes of the initial time points were used to compare the flux into the first step of each C4 pathway. Due to rapid incorporation of isotope in the wild type, additional time points were added for the early part of the labeling period. 13C incorporation into malate was more rapid in the wild type than in dct2-1, and in the wild type, enrichment leveled off within 60 s, suggesting that the singly labeled pool had achieved a maximum value (Figure 5B). In dct2-1, the labeling increased slowly and continued with time, indicating that malate accumulated in other subcellular locations (e.g., vacuoles) not intimately tied to assimilation, resulting in a larger overall pool size (Figure 5C).
13CO2 Labeling of Malate and Aspartate in Leaf Segments of 9-d-Old Wild-Type and dct2-1 Plants.
Leaf segments (sections B + C in Figure 1C) were labeled with 330 ppm 13CO2 at a light intensity of 500 µmol m−2 s−1 as previously described and analyzed for malate and aspartate labeling.
(A) Simplified metabolic network indicating the position of the singly labeled mass isotopomer [M+1]+ of malate and ASP in C4 photosynthesis. Other intermediates involved in the Calvin cycle can be more extensively labeled ([M+2]+ or [M+7]+) based on the number of carbon atoms. [M+i]+ is ith mass isotopomer. M is the base mass and i is the number of labeled carbons.
(B) Relative abundance (%) of [M+1]+ in malate and aspartate (sd, n = 3). Wild-type leaf segments were labeled for 1, 2, 5, 10, 20, 40, 60, 90, 120, 150, and 180 s . dct2-1 leaf segments were labeled for 20, 40, 60, 90, 120, 150, and 180 s. Solid (wild type) and dotted (dct2-1) lines indicate the linear rate of incorporation.
(C) Absolute pool sizes (nmol g FW−1) of malate and aspartate (se, n = 3) in leaf segments of wild-type and dct2-1 measured on LC-MS/MS. Samples were taken from unlabeled leaf segments.
(D) Initial incorporation rate of [M+1]+ carbon (nmol g FW−1 s−1) in malate and aspartate calculated from the pool sizes in (B) and the slopes in (C). Asterisk indicates that malate metabolism in dct2-1 was not determined (nd) from 13CO2 leaf labeling due to its accumulation in other places.
In wild-type plants, the rate of 13C incorporation into aspartate was significantly slower than into malate (2.9-fold; Figure 5D), consistent with published radiolabeling studies that described higher NADP-ME than PEPCK pathway activities (Hatch, 1971). Aspartate and malate labeling in the wild type plateaued within 60 to 90 s (Figure 5B), whereas these labeled metabolites in dct2-1 plants continued to increase at a much slower rate (Figure 5B). In the wild type, 13CO2 incorporation into the backbone of PEP through Rubisco-based assimilation resulted in aspartate mass isotopomers (i.e., [M+2]+ through [M+4]+; Supplemental Figure 2A). The higher masses were a measureable percentage of total labeling in the wild type, whereas aspartate remained mostly unlabeled or singly labeled in dct2-1 (Supplemental Figure 2B). In dct2-1, the 13C incorporation rate into aspartate was 1.1 nmol g FW−1 s−1 (Figure 5D), which is ∼6% of the wild type. This difference was consistent with the low photosynthetic rates measured in leaf sections B and C (3% or less of the wild type in dct2-1; Figures 1C and 1F). The changes were likely a consequence of the inability of dct2-1 to regenerate PEP, efficiently balance carbon and nitrogen across cell types, and maintain high rates of C4 metabolism, due to the disruption of the NADP-ME pathway. Previously, Wang et al. (2014b) suggested that the PEPCK pathway does not operate without other C4 pathways. Such a reliance on the PEPCK pathway would severely compromise photosynthesis and may partially explain the limited growth phenotype of dct2-1.
The contribution of the PEPCK pathway to carbon metabolism was further examined by combining the infrared gas analyses (IRGA; Figure 1F) with isotopic labeling rates of malate and aspartate (Figure 5D). The combined rate of assimilation in the wild type estimated from malate and aspartate isotope incorporation was 67.1 nmol gFW-1s−1 (Figure 5D). The dct2-1 rate of CO2 assimilation in leaf section C as determined by IRGA was 3% of the wild type (Figure 1F). For similar leaf thicknesses between lines, this would correspond to a total CO2 incorporation of 2.1 nmol gFW−1 s−1 (i.e., 3% of 67.1). Given that the aspartate label incorporation was measured to be 1.1 nmol gFW−1 s−1 (Figure 5D), the remainder (∼1.0 nmol gFW−1 s−1) comes from malate that was transported by a mechanism other than through DCT2 (consistent with Figure 3C) or by conversion to aspartate for import to the chloroplast as described below. Thus, PEPCK pathway activity is presumed to be enhanced in the dct2-1 line in a relative sense (∼55% of total net assimilation rate in dct2-1 versus 25% in the wild type). However, despite the apparent increase in PEPCK pathway activity, it is nonetheless insufficient to restore growth as dct2-1 PEPCK-based assimilation was ∼1.5% of the total wild-type assimilation rate (Figure 1F).
BS Cells Convert Malate to Aspartate and Alanine More Readily in dct2-1
The 13C experiments indicated that Calvin-Benson cycle activity was decreased in dct2-1 and reflected metabolic reprogramming. As the location and existence of certain C4 steps in the BS are unclear (Pick et al., 2011; Wang et al., 2014b), 14C studies with isolated BS cells were conducted to further probe the steps of C4 metabolism (Figure 6). The 14C studies indicated the routes of CO2 delivery to the chloroplast without requiring an intact C4 cycle. In wild-type BS cells, 14C-malate was taken up and converted to 14C-pyruvate, aspartate, and alanine (Figure 6A) and detected after separation on HPLC by scintillation counting. The separation of BS strands from M cells resulted in an incomplete NADP-ME cycle such that pyruvate accumulated following the decarboxylation of malate. Due to the deficiency of malate import in the dct2-1 plants, little pyruvate was detected ; however, two amino acids, aspartate and alanine, were more labeled in dct2-1 than the wild type (Figure 6A). Aspartate could serve as an alternative metabolite to transport carbon within the BS cells when the import of malate into the BS chloroplasts is disrupted. OAA and aspartate generated from malate may partially complement the malate transport deficiency and account for the production of labeled aspartate and alanine in dct2-1. In partial support of such a mechanism, mitochondrial and chloroplast-localized aspartate aminotransferases have been identified (Majeran et al., 2010), though the identification of an analogous alanine aminotransferases remains elusive.
14C-Malate, Aspartate, and Glutamate Labeling of BS Cells.
Wild-type (solid symbols) and dct2-1 (open symbols) plants were 14C labeled to describe BS photosynthetic operation (sd, n = 3). Inset shaded plots emphasize the initial 5-min incorporation.
(A) Time-course incorporation of 14C-malate into pyruvate, aspartate (ASP), and alanine (ALA).
(B) Time-course incorporation of 14C-ASP into PEP (square symbols) and ALA (red triangle symbols). Solid (wild-type) and dotted (dct2-1) lines indicated the linear incorporations after lag period.
(C) Time-course incorporation of 14C-glutamate into polar metabolites and 2-OG.
A comparable set of experiments that involved provision of 14C-aspartate to BS cells circumvented the requirement for an intact PEPCK pathway and were used to test the role of aspartate in providing carbon to the Calvin-Benson cycle. The linear rates of labeling in most cellular intermediates of the BS cells were similar in the wild type and dct2-1 (Supplemental Figure 3), with the exceptions of 3-PGA (Supplemental Figure 3D), PEP, and alanine (Figure 6B). PEP and 3-PGA were more rapidly labeled after a 5-min lag in the wild type, whereas labeling in alanine was greater in dct2-1. The increased radioactivity in 3-PGA and PEP reflected higher overall photosynthetic activities in the wild type, consistent with the net CO2 assimilation rate, 13C data, and differences in growth. The changes in nitrogen movement across the chloroplast envelope associated with aspartate use were likely balanced by alanine production and export and could account for the enhanced alanine labeling.
2H-Malate Labeling in BS Cells Supports Limited Mitochondrial Pathway Activity
2,3,3-2H-malate labeling experiments were used to further confirm aspects of aspartate movement in BS cells (Figure 7). 2H-malate was rapidly taken up and incorporated into fumarate (Figures 7A and 7B) in both the wild type and dct2-1. The steady state levels (∼15%) indicated the active pool of fumarate was quickly saturated. Both fumarate and OAA could be double labeled at different positions within each molecule and, as shown, would contribute to different combinations of single and double labeling in malate as well as aspartate (Figure 7A). Consistent with 14C-malate experiments, the provision of 2H-malate isotopically labeled alanine at low levels. The results support a mechanism where malate is used to generate OAA and aspartate in the mitochondria. The exchange between the organic acids would enable the removal of several deuteriums in this process, creating singly labeled aspartate that could be exported to the chloroplast and converted to alanine. Aspartate was ∼5 to 10% enriched by 5 min, whereas alanine was closer to 1%, though the labeling patterns were similar (Figure 7B). Singly labeled aspartate and alanine were more abundant than double-labeled mass isotopomers, suggesting 2H-malate is converted to fumarate or other intermediates prior to generation of the two amino acids. Similar to the 14C-labeling studies (Figures 6A and 6B), absolute 2,3,3-2H-malate incorporation into aspartate and alanine was higher in dct2-1 (Figure 7C). Other organic acids, including succinate and citrate, were also more labeled (Supplemental Figure 4A) in dct2-1, which suggests that mitochondrial involvement was enhanced relative to the wild type.
Time Course 2,3,3-2H-Malate Labeling of BS Cells in Wild-Type and dct2-1 Plants.
(A) Metabolites labeled from provision of 2,3,3-2H-malate (purple box) in BS cells. [M]+ is the unlabeled mass isotopomer. [M+1]+, [M+2]+, and [M+3]+ contain one to three deuterium atoms. Orange and blue lines indicate reactions that could lead to [M+1]+ and [M+2]+ mass isotopomers, respectively, of alanine and aspartate. Only directly impacted heteroatoms (i.e., hydrogens) are presented.
(B) Labeling pattern of fumarate (FUM), aspartate, and alanine in the wild type (black/gray) and dct2-1 (red/dotted). All data were corrected for natural isotope abundance (sd, n = 4).
(C) Quantification of deuterium isotope incorporation (nmol gFW−1) into aspartate and alanine from labeled malate (sd, n = 4). Data were calculated from pool sizes of aspartate and alanine in BS cells and the summed relative labeling data in (B). Solid (wild type) and dotted (dct2-1) lines reflected the linear incorporation rates that were quantified as slopes (nmol gFW−1 s−1).
Conversion of Glutamate to 2-Oxoglutarate Is Unaffected by dct2-1 Mutation
In Arabidopsis thaliana, dicarboxylate transporters OMT1 and DCT1 were shown to be essential for photorespiration (Kinoshita et al., 2011); At-DCT2 (the ortholog of Zm-DCT2), however, is not expressed in photosynthetic leaf tissue in Arabidopsis (Renné et al., 2003). To examine the potential involvement of DCT2 in photorespiration in maize, we measured photorespiratory pool sizes and quantified isotopic labeling from 13CO2 in dct2-1 and wild-type plants (Supplemental Figure 5). Although the 2-phosphoglycolate (2-PG) pool size was reduced in dct2-1, glycine and serine concentrations were elevated and less labeled. The reduced 2-PG pool and increased amino acid concentrations do not support a limitation in nitrogen substrate availability for the related reactions (Supplemental Figure 5A).
Photorespiratory links with malate transport and nitrogen metabolism were further probed by decoupling the pathway and providing the source of nitrogen exogenously. 14C-glutamate was supplied to isolated BS cells and incorporation of label into polar metabolites was quantified. As shown in Figure 6C, the linear incorporation of 14C into polar metabolites including 2-OG was similar or slightly less in dct2-1 relative to the wild type and suggested that nitrogen exchange in BS cells was not significantly compromised by the mutation. Although the malate availability for counter-port activity could be compromised in dct2-1 plants, alternative reactions catalyzed by glutamine or asparagine synthetase, glutamate synthase, and glutamate dehydrogenase outside of the chloroplast could also be involved in ammonia recycling (Becker et al., 2000; Miflin and Habash, 2002; Valadier et al., 2008). Transcriptionally upregulated asparagine synthetase (Supplemental Data Set 1) and the accumulation of asparagine (Supplemental Table 1) were observed in dct2-1.
Transcriptome Analysis Indicates Extensive Transcriptional Response to the Loss of DCT2
RNA-seq analysis was performed to inspect global gene expression changes in dct2-1 plants. The mRNA profiles of mutant and wild-type individuals were assessed at the −4, −1, +4, and +10 leaf developmental stages (Figure 1C). The difference in gene expression in dct2-1 compared with the wild type was calculated for each gene as the log2 (dct2-1 fragments per kilobase of transcript per million mapped reads [FPKM]/wild-type FPKM), using CuffDiff (see Methods), with a false discovery rate of 0.05. Of the 30,634 genes expressed in the leaf tissue, 13,439 (44%) showed a significant difference in expression for at least one of the tested segments (Supplemental Table 3) and were subject to further analysis. To identify metabolic pathways that were affected at the transcriptional level, gene IDs were assigned to pathways using the MapMan pathway annotation for maize (Thimm et al., 2004; Usadel et al., 2009).
Many pathways, including photosynthesis, cell wall metabolism, transport, hormone, signaling, transcriptional regulation, and lipid metabolism, were affected in the mutant plants (Supplemental Table 3). In agreement with the labeling data, photosynthesis genes, such as photosystem I and II; transport genes, including DCT1 and OMT1; Calvin cycle genes; and other C4 genes, including NADP-ME and PEPCK pathways, showed lower transcript levels (Figures 8A to 8D; Supplemental Table 3). In addition, the levels of transcripts encoding chlorophyll biosynthesis proteins were reduced, reflecting the pale green leaf phenotype of dct2-1 (Figure 8E; Supplemental Data Set 1). Starch synthesis transcripts also appeared to be lower and starch degradation increased in the mutants, explaining the lack of starch granules in dct2-1 leaves (Figures 8F and 8G; Supplemental Data Set 1). Transcripts for genes related to photorespiration were either slightly reduced or unchanged in their expression pattern (Figure 8H; Supplemental Data Set 1). Transcripts for an aspartate aminotransferase gene (GRMZM2G094712), predicted to be localized in mitochondria (Majeran et al., 2010), and two alanine aminotransferase genes (GRMZM5G828630 and GRMZM2G088064) accumulated to higher levels in dct2-1, consistent with the higher levels of aspartate and alanine identified by the isotopic labeling experiments. To date, amino acid transporters in plants remain largely uncharacterized, though the labeling results suggest a heightened need for such transporters during C4 photosynthesis in dct2-1. Two more genes that showed elevated transcript levels in the mutants may be candidates to move aspartate and alanine during C4 photosynthesis: GRMZM2G042933, a putative amino acid transporter predicted to be localized in the chloroplast (Majeran et al., 2010), and GRMZM2G149619, a putative bile acid sodium symporter family protein predicted to be localized in the chloroplast (Majeran et al., 2010) (Supplemental Table 3).
Heat Map Representation of the Differences in Gene Expression between dct2-1 and Wild-Type Plants.
RNA-seq was performed on four segments along the developmental gradient of 9-d-old plants. Values represent log2 (dct2-1 FPKM/wild-type FPKM). Red indicates upregulation in dct2-1 relative to the wild type, and green indicates downregulation in dct2-1 relative to the wild type. Each heat map is accompanied by a significance plot: black squares indicate a statistically significant difference in gene expression in dct2-1 relative to the wild type in a specific segment; gray squares represent unsignificant differences. Squares in each row, moving from left to right, represent segments −4, −1, +4, and +10 (as defined in Figure 1C).
Several large transcription factor families showed significant changes in expression; in some cases, the differences between dct2-1 and the wild type exceeded 100-fold. For example, HB (homeobox) transcription factors, known to be involved in cell differentiation, control of cell growth, and patterning, showed large changes in all segments, as did GATA transcription factors that are known to be involved in light signaling (Supplemental Data Set 1). Though beyond the scope of this report, the transcriptomic data suggest several lines of further study to characterize regulatory factors that may be responsive to altered C4 metabolism.
DISCUSSION
C4 metabolism involves the combination of up to three photosynthetic carbon assimilatory pathways across two cell types, requiring coordination of enzymatic and transport activities across multiple organelles. This level of complexity has hindered the interpretation of many labeling studies and limited understanding of this crucial metabolic pathway in plants. Recent depictions of C4 photosynthesis have recognized that the individual subtype pathways are complementary in function (Furbank, 2011; Pick et al., 2011). For example, the PEPCK pathway can move nitrogen as well as carbon, the NADP-ME pathway produces reducing equivalents in the BS chloroplast, and NAD-ME produces reducing equivalents in the mitochondria that may be further used to produce ATP. In this study, multiple stable and radioisotopic labeling experiments, enzyme assays, transcriptome studies, and morphological data were used to conduct a mechanistic study of C4 pathway use in wild-type maize and to examine the metabolic consequences of the disruption of the primary malate carbon shuttle pathway in a transposon-tagged dct2-1 mutant plant line. The use of 13C-labeling, transcript profiling, and microscopy permitted a detailed in planta characterization of activities, whereas studies of isolated BS cells, BS chloroplasts, and enzyme assays enabled a more thorough metabolic characterization of activities. Though the latter can be affected by changes in concentration or be potentially biased by purity of cellular isolation or intact state of chloroplasts, our analyses were based on comparative studies; therefore, any artifacts would similarly affect the wild type as well as the mutant lines. Furthermore, the interpretation was supported by measurements that judged, for example, organelle quality prior to further evaluation. The investigations articulate the concerted use of NADP-ME and PEPCK pathways that are altered in the dct2-1 line and contribute to a more comprehensive understanding of C4 photosynthesis.
DCT2 Is a Primary Transporter for Malate into the BS Chloroplast during C4 Photosynthesis
Higher plant genomes contain paralogous groups of genes that perform similar functions. Many of the proteins thought to be involved in C4 photosynthesis are members of such paralogous groups (Taniguchi et al., 2004; Rondeau et al., 2005; Shenton et al., 2006; Studer et al., 2014). Although this redundancy likely enables the evolution of new activities, it also makes it difficult to determine which of the genes have been co-opted to participate in C4 photosynthesis (Bräutigam et al., 2008; Majeran and van Wijk, 2009; Weber and von Caemmerer, 2010). Three paralogous dicarboxylate transporters that can transport malate at high efficiency have been characterized in maize leaves: DCT2, DCT1, and OMT1 (Taniguchi et al., 2004). OMT1 is differentially expressed at high levels in maize M cells (Li et al., 2010; Tausta et al., 2014) and has been hypothesized to transport OAA into the M chloroplast in exchange for malate (Bräutigam et al., 2008). DCT1 is expressed at low levels and was shown to be enriched in the M in maize. DCT1 is also expressed in the BS (Li et al., 2010; Chang et al., 2012; Tausta et al., 2014), but the precise metabolic function has yet to be determined. DCT2 is a candidate for malate import into maize BS chloroplasts because it is highly and differentially expressed in BS cells with a gradated expression along the developing leaf and transcript and protein levels that are light responsive (Taniguchi et al., 2004; Bräutigam et al., 2008; Li et al., 2010; Tausta et al., 2014). In this study, a combination of physiological, biochemical, transcriptional, and morphological data were combined to elucidate the function of dct2-1 in maize leaves. The data strongly suggest that DCT2 enables malate flux into the BS chloroplast during C4 photosynthesis.
The Loss of DCT2 Hinders the Photosynthetic Development of dct2-1 Plants
The loss of DCT2 function reduces photosynthesis and resulted in extensive changes in gene expression. Over 44% of all expressed genes spanning most major metabolic pathways showed a significant change in their expression level (Supplemental Data Set 1; Figure 8). Transcription factor abundance was also altered (Supplemental Data Set 1) and indicated transcriptional reprograming in response to the loss of DCT2. The impaired photosynthetic activity of the mutant was additionally evident from the reduced assimilation of CO2 and reduced expression of associated photosynthetic genes (Figures 1F and 8A) as well as changes to starch accumulation apparent from the electron microscopy images (Figure 2) and genes encoding starch biosynthesis (Figures 8F and 8G; Supplemental Table 3). The absence of DCT2 hindered the photosynthetic developmental gradient, altering chlorophyll levels and net CO2 assimilation rates along the longitudinal axis of maize leaves. TEM analysis also indicated that thylakoid membranes from chloroplasts in BS and M cells were developmentally compromised in dct2-1 leaves (Figure 2). Despite these deficiencies, seedlings were viable and capable of supporting low levels of growth at 1 month of age (Figure 1D).
Both the NADP-ME and PEPCK Pathways Participate in C4 Photosynthesis in the Wild Type and dct2
13C labeling studies indicated that the NADP-ME pathway is 2.9-fold more active than the PEPCK pathway in the wild type (Figure 5D), consistent with a prior report (Hatch, 1971). Labeling experiments with 14C-malate performed on isolated BS strands indicated that the DCT2 mutation limits the operation of the NADP-ME pathway in the BS and negatively impacts the PEPCK pathway. Since the 14C-ASP incorporation rates into most cellular intermediates of BS cells were similar in both the wild type and dct2-1, the limitation in the PEPCK pathway likely involved the regeneration of PEP, which is shared by all C4 subtype pathways. The operation of a PEPCK pathway is insufficient to maintain robust growth in the absence of a NADP-ME pathway, as suggested by Wang et al. (2014b). Calculations based on results presented here suggest that PEPCK pathway activity may be elevated to closer to one-half of all assimilatory activity in the mutant; however, the mutant’s growth rate was slow, as indicated by 13CO2 leaf labeling, and the net assimilation was only 3% of the wild type based on results from IRGA.
Alanine Production in dct2-1 May Facilitate PEP Regeneration and Balanced Nitrogen Metabolism
One of the more intriguing observations from 14C labeling was the generation of quantifiable 14C in two amino acids, 14C-ASP and 14C-ALA (Figures 6A and 6B). 2,3,3-2H-malate labeling studies further confirmed these findings and supported the involvement of mitochondria reactions in maize C4 metabolism (Figure 7). The rate of production of these two amino acids was greater in the dct2-1 plants and may be necessary to sustain the regeneration of PEP for continued C4 operation. Balancing the export of alanine to match elevated aspartate import in multiple locations would facilitate the movement of both carbon and nitrogen within C4 metabolism (Pick et al., 2011) (Figure 9) and is the simplest interpretation of the combined experiments. As one possibility, some aspartate derived from malate in the mitochondria could be imported into the chloroplast and used to make low levels of labeled alanine that were not evident until close to 5 min (Figure 7B). The conversion of malate to OAA in the mitochondria would produce NADH that could be further used to generate ATP by oxidative phosphorylation. Greater ATP production and possibly enhanced respiration (Figure 1F) could be necessary to support the observed enhancement in PEPCK pathway activity in dct2-1. The transcriptome data are consistent with these ideas, as one aspartate aminotransferase (GRMZM2G094712) predicted to be localized in the mitochondria and two alanine aminotransferases (GRMZM5G828630 and GRMZM2G050481) with unknown targeting accumulated to higher levels in dct2-1. Chloroplast-targeted aspartate aminotransferases have also been previously described (Majeran et al., 2010). Thus, the genes involved in the metabolism of these amino acids may be transcriptionally altered in response to increased levels of unmetabolized malate. In addition, labeled alanine was observed in BS chloroplasts obtained from the BS cells labeled with 2H-malate, though pooling of these samples did not allow for technical replicates (Supplemental Figure 4B). Thus, metabolism of malate by alternate mechanisms may be of greater importance in dct2-1 carbon assimilation.
C4 Photosynthesis Including Coordination between Subtype Pathways.
In maize, C4 photosynthesis involves the movement of carbon between M and BS cells using the NADP-ME and, to a lesser extent, the PEPCK pathways (black lines). Mitochondrial reactions may play a limited role in both wild-type and dct2-1 lines. Blue lines indicate steps that can partially compensate for reduced malate transport in the dct2-1 line. The provision of aspartate results in enhanced malate import in the wild type where the aspartate is either recycled directly or used to produce nitrogen and OAA. OAA could be exported or contribute to CO2 generation and the production of alanine that is more prominent in the compromised dct2-1 line. Dashed lines indicate reactions for which more information is needed.
Photorespiration in dct2-1 Plants
Photorespiration in C4 plants is greatly reduced compared with C3 species but remains important for detoxification of 2-PG (Zelitch et al., 2009) to distribute reducing equivalents through multiple subcellular organelles and to produce glycine, serine, and folate that may be used elsewhere (Sweetlove et al., 2006; Collakova et al., 2008). Additionally, CO2 production through photorespiration may help sustain Calvin-Benson cycle activity (albeit less efficiently) when CO2 concentrations are reduced (Igamberdiev, 2015). Compromised DCT2 operation and concomitant low flux through serine and glycine were nevertheless sufficient for the dct2-1 plants to prevent 2-PG accumulation and perform photorespiration necessary to parallel other reductions in Calvin-Benson cycle metabolism and growth. Metabolism was analyzed in dct2-1 lines with 14C- and 13C-labeling and metabolite pool measurements. The pool sizes for serine and glycine were elevated in dct2-1, whereas 2-PG, the first intermediate of the pathway, was reduced. Labeling in amino acids and sugar phosphates not limited to 2-PG, serine, or glycine (Supplemental Figure 5B) were substantially reduced in dct2-1, indicating that the effects were not specific to photorespiration. That 2-PG was clearly labeled in dct2-1 (17% ± 4% average 13C enrichment by 3 min; Supplemental Data Set 2) further suggested some functional pathway operation. Though changes in flux cannot be inferred from pool size information alone, the observed decrease in 2-PG concentration and increase in serine and glycine pools indicated a rebalancing of metabolism that would not suggest a limitation in photorespiration or nitrogen exchange in dct2-1.
Next, the provision of nitrogen for glycine biosynthesis was decoupled from the regeneration of glutamate and malate cotransport by providing 14C-glutamate to isolated BS cells. A time course of radioactive isotope incorporation indicated that the transient labeling into 2-OG was comparable in the wild type and dct2-1 (Figure 6C). Though conversion of glutamate to 2-OG through aminotransferase reactions is not limited to photorespiratory activity, in isolated BS cells that have a primary role in photosynthesis, other uses for nitrogen such as protein biosynthesis are less likely to be significant within the time frame of labeling. Prior work has confirmed that the production and turnover of most amino acids in leaves occurs at rates far lower than those associated with Calvin-Benson cycle activity (Ma et al., 2014).
Mechanism of Malate Transport and Coordination with Aspartate
In this study, C4 photosynthesis with compromised malate transport was examined by a combination of experiments with isolated BS chloroplasts from wild-type and dct2-1 leaves. In measurements based on the formation of pyruvate, malate transport and conversion to pyruvate were enhanced in the presence of aspartate, which has also been observed in wild-type maize by others, though without a conclusive explanation (Chapman and Hatch, 1979; Jenkins and Boag, 1985; Kanai and Edwards, 1999). It is possible that aspartate taken up by a separate mechanism serves in an antiport capacity with malate as it has been suggested that aspartate is not metabolized in the process (Boag and Jenkins, 1985). However, in prior studies, the role for aspartate was evaluated based on tracing 14C-aspartate movement in isolated chloroplasts and may not fully capture modifications to aspartate that were unrelated to the carbon backbone or account for observations we made specific to dct2-1.
A more involved scheme is proposed in Figure 9 where import of aspartate could be recycled directly with malate or used to produce OAA through a putative aspartate aminotransferase targeted to the chloroplast (Majeran et al., 2010). In wild-type maize, aspartate could serve in a recycling capacity (no net movement) to drive malate import and also provide a source for the PEPCK pathway. Some of the imported aspartate could also be converted and exported as OAA, which is a known antiport counter substrate for malate by DCT2 in maize (Taniguchi et al., 2004). Alternatively, the OAA could be subsequently metabolized to malate for decarboxylation prior to generating alanine that would balance the movement of nitrogen. Our data further support some conversion of malate to aspartate that is enhanced in dct2-1 (Figure 6) and may possibly occur in the mitochondria prior to movement into the chloroplast (Figure 7). This would provide an alternative use of malate that is greater in the mutant where it may be more crucial due to the compromised primary malate shuttle pathway (Figure 9). Excess nitrogen could be returned to the cytosol through conversion of pyruvate to alanine and export of this amino acid, consistent with enhanced aspartate and alanine labeling in dct2-1 by multiple experiments (Figures 6 and 7), including alanine labeling in chloroplasts isolated from labeled BS cells (Supplemental Figure 4B). In this model, pyruvate that results from decarboxylation of directly imported malate by NADP-ME in the wild type would be exported from the BS chloroplasts without comovement of ammonia. Pyruvate import into the M cell plastids in C4 plants could involve Na+ or H+ (Flugge et al., 1985; Ohnishi and Kanai, 1987a, 1987b). In maize, Ohnishi et al. (1990) suggest the driving force is a H+ gradient, as reviewed by Heldt (2002). The presence of a pyruvate transporter in both M and BS cells was described (Furumoto et al., 2011), though the mechanism of pyruvate export from BS plastids remains unknown (Weber and von Caemmerer, 2010). The model presented also infers the presence of a BS-plastid-localized alanine aminotransferase, and although expression was elevated in dct2-1, the location to the BS plastids remains speculative. An attractive feature of the model is that it provides an additional mechanism to balance carbon and nitrogen metabolism across multiple cellular locations. The lack of response to aspartate in enzyme activity assays of dct2-1 isolated chloroplasts (Figure 3) could reflect the fact that the antiport activity is unnecessary when malate import is compromised. In this case, any limited import of aspartate could be metabolized to form alanine, balance nitrogen, and provide additional CO2, though this would alter production of reducing equivalents and thus not necessarily be reflected in oxidation measurement of the enzyme assay.
METHODS
Ac Reverse Genetic Screening
Mutagenesis of the DCT2 locus was performed using bti99221::Ac, located 401.2 kb (0.44 cM) 5′ of the gene, as a donor Ac. Populations for identifying Ac insertions into DCT2 were generated as described by Singh et al. (2003). One thousand and fifty high-dose kernels were planted in 11.0 × 21.5-in 50-cell hexagon plug trays (TO Plastics) filled with Metro-360 soil (Sun-Grow) and turface at a ratio of 3:1 by volume. Flats were grown in a greenhouse with 16 h light per day, with supplemental light of 500 μmol m−2 s−1, 28°C, and 40% humidity. A one-eighth-inch punch of tissue was sampled from each plant 10 d after sowing. Samples were pooled in groups of 10 into a single tube. DNA was extracted using the cetyltrimethyl-ammonium bromide protocol for DNA isolation: Leaves were ground in liquid nitrogen and incubated at 65°C for 1 h in extraction buffer (0.35 M sorbitol, 0.15 M Tris, 5 mM EDTA, 2 M NaCl, 0.83% cetyltrimethyl-ammonium bromide, and 0.83% N-lawoylsarcosinl). To each tube an equal volume of chloroform-isoamyl alcohol (24:1) was added, and the samples were mixed well and centrifuged at 12,000g for 10 min at 4°C. The supernatant was transferred to a new tube containing isopropanol. Samples were then centrifuged at 12,000g for 15 min at 4°C, the liquid was discarded, and the DNA pellet was washed with 70% ethanol. The samples were then centrifuged at 12,000g for 3 min at room temperature and air dried, and then the pellet was dissolved in 10 mM Tris-HCl, pH 8. Six primer sets were designed to cover a 16-kb region containing DCT2 and 2 kb upstream and downstream of the gene. Each genomic primer was paired with two Ac-specific primers (one for the Ac 5′ and one for the Ac 3′). Primer sequences are listed in Supplemental Table 2. Amplification was done using Phire Taq (Thermo Scientific F-122L) according to the manufacturer’s recommendations. PCR products were separated on a 1% agarose gel by electrophoresis. Pools containing putative insertions were deconvoluted to a single plant using the same DNA extraction and PCR amplification procedures described above. Confirmed insertions were mapped using Sanger sequencing of the amplified insertion product using the Ds end primer.
qPCR of 9-d-Old Maize Leaves
Whole leaf samples (two biological replicates) from wild-type and mutant plants were each ground in liquid nitrogen. Total RNA was extracted using Trizol reagent (Invitrogen) according to the manufacturer’s instructions. RNA samples were treated using DNase I recombinant, RNase-free (Roche) to remove any contaminating DNA. cDNA was synthesized from each DNase-treated RNA sample using a Transcriptor First Strand cDNA Synthesis Kit (Roche) with Anchored Oligo (dT)18 primers. Two cDNA preps were performed for each sample along with a prep lacking the reverse transcriptase. Quantification of target gene expression was calculated relative to the calibrator gene using advanced relative quantification in the LightCycler 480 SW 1.5 software (Roche). In each segment, the ratio of the expression level of the calibrator gene to the target gene was calculated using the software. Reactions were run with forward primer 5′-TTCTTGCAGTCTCACTATGGAT-3′ and reverse primer 5′-GTTTAGCACTCCAAGCACA-3′ using LightCycler 480 SYBR Green I Master (Roche) in a Roche LightCycler 480 II real-time PCR Instrument. The reaction profile was as follows: 95°C for 5 min, 45 cycles of 10 s at 95°C, 10 s at 60°C, and 10 s at 17°C, followed by one cycle of 95°C for 5 s and 65°C for 1 min; samples were left at 95°C without cooling. Three technical replicates of each sample were run.
RNA-Seq Analysis of the Maize Leaf Transcriptome
One-centimeter leaf segments (−4, −1, +4, and +10 in Figure 1C) from 9-d-old plants, the third leaf on wild-type and mutant plants, were frozen in liquid nitrogen. Samples were pooled from 15 plants per biological replicate (three replicates for each segment of each wild type and mutant). Total RNA was extracted using TRIzol (Invitrogen) according to the manufacturer’s protocol and treated with DNase prior to library construction. RNA-seq libraries were constructed according to the protocol described by Wang et al. (2014a). Libraries were sequenced on the Illumina HiSequation 2000 machine. The number of raw reads obtained for all RNA-seq libraries is listed in Supplemental Table 3. Reads were sequenced, deconvoluted, and filtered using the manufacturer’s default pipeline and parameters. The reads were aligned to the maize reference genome B73 AGPv2 using GSNAP (Genomic Short-read Nucleotide Alignment Program) (Wu and Nacu, 2010). Differentially expressed genes between mutant and wild-type plants were identified using CuffDiff (Trapnell et al., 2010) with 0.05 as the allowed false discovery rate. All raw RNA-seq data were deposited in NCBI under accession number GSE67722. Heat maps were generated using the R package ggplot2 (Ginestet, 2011).
Measurements of Net Photosynthetic Rate and Chlorophyll Contents
An infrared gas analyzer, the LI-6400 XT portable photosynthesis system (Li-Cor), was used to measure the net photosynthetic rates of wild-type and mutant leaves at midday, under a light intensity of 500 µmol m−2 s−1 (MP 400W/H75/UVS/PS/740 lamps; Venture Lighting). Chlorophyll contents in 10 mg of fresh wild-type and mutant leaf tissues at midday were extracted with 10 mL 80% (v/v) acetone and determined spectrophotometrically according to Arnon (1949).
TEM of Leaf Sections
Two-millimeter cross-sectional strips of leaves from 9-d-old wild-type and dct2-1 plants were fixed for 2 h in 2% gluteraldehyde in 100 mM, pH 6.8, PIPES buffer at room temperature. The leaves were then washed three times in 100 mM, pH 6.8, PIPES buffer. The leaves were fixed in buffered osmium tetraoxide for 1.5 h and rinsed three times in water. The leaves were dehydrated in an ethanol/acetone series as follows: 5, 10, 20, 30, 50, 75, and 95% ethanol for 20 min each, followed by 30 min in 100% ethanol, 15 min in 100% acetone, and a second 45-min incubation in fresh 100% acetone. The leaves were infiltrated with Spur’s resin (Cat. No. RT14300; Electron Microscopy Science) dissolved in 100% acetone as follows: 12 h, 5%; 12 h, 10%; 24 h, 25%; 24 h, 50%; 24, 75%; and 24 h, 100%. The leaves were then embedded in 100% resin and incubated at 60°C for 2 d. The resin blocks were cut into 0.1-µm sections using a Leica Ultracut UCT microtome. The sections were stained in uranyl and lead salts. Digital images were acquired using a LEO 912 AB energy filter transmission electron microscope operated at 120 kV. In some cases, large fields of view were acquired by montaging.
13C Labeling and Mass Spectrometry Analysis of Metabolites
Leaf segments (sections B and C in Figure 1C) of 9-d-old wild-type and dct2-1 plants were labeled in custom-made individual gas-tight chambers by blowing through premixed gas containing 13CO2 (13CO2/N2/O2 ratio of 0.033/78/21.967) at a flow rate of 2 L/min and a fluorescent light intensity of 500 µmol m−2 s−1 at 28°C. Labeled samples were collected over a 3-min interval at 1, 2, 5, 10, 20, 40, 60, 90, 120, 150, and 180 s for the wild type and 20, 40, 60, 90, 120, 150, and 180 s for dct2-1 plants followed by immediate quenching with liquid nitrogen. Samples were extracted and analyzed on LC-MS/MS and GC-MS for labeling of metabolites according to Ma et al. (2014).
Preparation of Maize BS Cells
Bundle sheath cells were isolated from the third leaf of 10- to 12-d-old plants. Leaf biomass (∼2 g of fresh weight) was homogenized in an isolation buffer containing 350 mM sorbitol, 50 mM Tris-HCl, pH 8.0, 5 mM EDTA, 0.02% BSA, and 10 mM MgCl2 using a PowerGen 500 homogenizer (Fisher Scientific) for 1 min at maximum speed. BS cells were collected and separated from M cells by passing through 40 mesh nylon cloth and then washed two additional times with isolation buffer to remove mesophyll cellular debris. Cell purity was confirmed by light microscopy. Harvested BS cells were maintained in isolation buffer until use within <10 min.
14C Labeling and Processing of Metabolites from Isolated BS Cells
BS cells (50 mg of fresh weight) were placed in individual wells of a 48-deep well plate with 0.5 mL isolation buffer. Cells were preincubated on a rocking platform exposed to a fluorescent light intensity of 500 µmol m−2 s−1 for 10 min. Then, 0.5 μCi of either uniformly labeled malate, glutamate, or aspartate was added to 50 μL isolation buffer for durations including 0.25, 1, 2, 5, 10, 30, and 60 min with constant shaking. Reactions were stopped by the addition of cold chloroform/methanol (3/7, v/v). Polar and nonpolar phases were separated by the addition of water (final concentration 44%) and centrifugation at 2200g for 10 min at 4°C. The top (polar layer) was removed. Water extraction was repeated two times and aliquots were combined. The entire experiment was repeated three times. To ensure the quenching was adequate, the chloroform/methanol was added in a control experiment prior to the addition of cells, substrate, and isolation buffer and then incubated for the maximum length of the experiment (1 h), but did not result in labeled metabolites.
Amino acids were separated from anionic and neutral fractions by cation exchange. The amino acids that are bound to the column were eluted with 1 M NH4OH, prior to drying with nitrogen aspiration at 45°C.
Identification and quantification of labeled products was performed using HPLC with in-line scintillation counting. Compounds were separated using a Waters 1525 HPLC with binary pumps with a 2707 autosampler set at the partial loop mode to assure all radiolabel was injected. Radiolabeled peaks were detected with in-line scintillation counting using a Perkin-Elmer 150TR flow scintillation analyzer with Ultima-Flow M (Perkin-Elmer) scintillation fluid pumped at a ratio of 1:4 (one part of radiolabeled sample to four parts of scintillation fluid). Conversion of peak area to total DPM injected was accomplished with the development of a standard curve of a range of known labeled standards.
Anionic metabolic intermediates that were not retained by the cation exchange were separated by reversed phase HPLC using a Synergi Hydro-RP 4u 80A, 150 × 2.00-mm column (Phenomenex) equilibrated with buffer A (10 mM TBA and 15 mM acetic acid, pH 4.5) and eluted with buffer B (100% methanol), with the following gradient: 0 to 5 min, 0% B; 5 to 50 min, 45% B; and 50 to 52 min, 90% B; at a flow rate of 0.3 mL/min.
Cationic compounds including amino acids were separated by hydrophilic interaction liquid chromatography HPLC using an EMD ZIC-cHILIC 3-µm 150 × 2.1-mm column (Merck) equilibrated with buffer B (95% acetonitrile and 0.1% formic acid) and eluted with buffer A (10 mM ammonium formate and 0.1% formic acid), with the following gradient: 0 to 5 min, 0% A; 5 to 60 min, 12.875% A; and 60 to 80 min, 45% A; at a flow rate of 0.5 mL/min.
The metabolites were identified by retention times that were first established using LC-MS/MS with unlabeled samples and the same column and gradient. Spiking experiments were used to judge adequate recovery and losses due to processing. Recovery of total counts in the presence of biological matrix or media only were as follows; glutamate, 89% and 98%; malate, 69 and 80%; aspartate 97% and 110%, respectively.
Isolation of BS Chloroplasts
BS chloroplasts were isolated by the method of Jenkins and Boag (1985). About 10 g of young maize leaves were deribbed and then sliced transversely with a razor blade. Leaves were immersed in 100 mL of precooled blending medium (0.33 M sorbitol, 10 mM HEPES-KOH, pH 7.8, and 5 mM EDTA) in a tall 300-mL beaker. Leaves were blended two times with a Polytron homogenizer (PT-MR 3100; Kinematica) for 60 s. The solution was filtrated through two layers of Miracloth and washed twice with an excess volume of the blending medium. The blending and wash procedures were repeated an additional two times. The BS cells were suspended with 80 mL of digestion medium (0.33 M sorbitol, 10 mM MES-KOH, pH 5.5, 1 mM K-Pi, 0.1% BSA, 2% Sumizyme C [Shin Nihon Chemical], and 0.1% Pectolyase Y-23 [Kyowa Chemical Products]) and incubated at 30°C for 20 min in the dark. At the end of the incubation period, the cells were lightly washed with blending medium and then agitated in 80 mL of chloroplast isolation medium (CIM; 0.33 M sorbitol, 25 mM HEPES-KOH, pH 7.8, 5 mM EDTA, and 0.1% BSA) to release chloroplasts into the medium. The solution was filtrated through 355-, 75-, 39-, and 20-µm nylon sieves in sequence, and the filtrate was centrifuged at 400g for 3 min at 4°C. The pellet of crude chloroplasts was suspended in 4 mL of CIM. Each 2 mL of the chloroplast suspension was layered on top of 4 mL of CIM containing 20% Percoll (GE Healthcare) and centrifuged at 700g for 3 min. The pellets were resuspended in 4 mL of CIM without BSA and the suspension was centrifuged again. The chloroplasts were resuspended in several hundred microliters of CIM. Chlorophyll concentrations were determined in 96% ethanol (Wintermans and De Mots, 1965). The intactness of the chloroplast preparations was estimated by measuring NADP-ME activities before and after the addition of 0.05% Triton X-100. NADP-ME was assayed spectrophotometrically with a reaction medium containing 50 mM HEPES-KOH (pH 8.0), 20 mM MgCl2, 2.5 mM EDTA, 0.5 mM NADP+, 5 mM DTT, and 5 mM malate. Intact state, quantity, and purity of chloroplasts were investigated by light microscopy (BX51; Olympus). Plastid number was evaluated using a hemocytometer.
Malate-Dependent Pyruvate Formation of BS Chloroplasts
The assay for pyruvate formation was performed as described by Jenkins and Boag (Boag and Jenkins, 1985, 1986; Jenkins and Boag, 1985). The reaction mixture contained 0.35 M sorbitol, 25 mM Tricine-KOH (pH 8.0), 2 mM MgCl2, 1 mM K-Pi, 10 mM KHCO3, 1 mM ADP, 5 mM 3-phosphoglycerate, 1000 units mL−1 catalase, 10 mM malate, and in some cases 10 mM aspartate. In the case of lysed chloroplasts, 0.5 mM NADP+ and 0.05% Triton X-100 were supplemented to the reaction mixture. After preincubation at 30°C (1000 µmol m−2 s−1), reactions were started by the addition of BS chloroplasts (5 µg chlorophyll mL−1). The reactions were quenched by the addition of HClO4 (final concentration 4.6%) and neutralized with 5 M KOH. After centrifugation, pyruvate in the supernatant was determined by spectrophotometrical measurement of NADH oxidation with assay mixture containing 0.3 M triethanolamine-HCl (pH 7.6), 3 mM EDTA, 0.1 mM NADH, and 2 units mL−1 lactate dehydrogenase.
2,3,3-2H-Malate Labeling of Isolated BS Cells and Chloroplasts
BS cells (50 mg FW) were preincubated with 0.5 mL isolation buffer in a 48-deep well plate for 10 min at a fluorescent light intensity of 500 µmol m−2 s−1 and 28°C. Fifty microliters of 2,3,3-2H-malate (10 mM) was added to start the time-course labeling for durations including 0, 5, 15, 45, 90, 120, 180, 300, and 3600 s. This time course was chosen based on the 14C studies in which the linear incorporation usually lasted for 5 min. Metabolism was then quenched by addition of 1.3 mL chilled chloroform/ methanol (3:7, v/v). Polar and nonpolar phases were separated by the addition of water. Polar metabolites (organic acids and amino acids mainly) were analyzed on LC-MS/MS and GC-MS as previously described.
BS chloroplasts were generated from 2 g (FW) BS cells and incubated with 0.5 mL isolation buffer at 500 µmol m−2 s−1 and 28°C for 10 min. 2,3,3-2H-malate (1 µmol) was added to start the labeling. After 1 h, labeled chloroplasts were centrifuged at 900g at 4°C for 10 min and decanted. The chloroplast pellets were resuspended in 2 mL isolation buffer and centrifuged again to remove labeled substrate. The rinse process was repeated three times to remove residual 2,3,3-2H-malate. Polar metabolites in chloroplasts were extracted with chloroform/ methanol/water and analyzed by LC-MS/MS and GC-MS.
Accession Numbers
Sequence data from this article can be found in the GenBank/EMBL databases under the following accession numbers: GRMZM2G040933 (DCT1), GRMZM2G086258 (DCT2), and GRMZM2G383088 (OMT1).
Supplemental Data
Supplemental Figure 1. 13CO2 labeling of wild-type and dct2-1 plants.
Supplemental Figure 2. Fractional abundances (%) of aspartate mass isotopomers in 13CO2-labeled leaf segments of 9-d-old wild type and dct2-1.
Supplemental Figure 3. Time-course 14C-Asp labeling of BS cells.
Supplemental Figure 4. 2,3,3-2H-malate labeling of BS cells and BS chloroplast in 9-d-old wild-type and dct2-1 plants.
Supplemental Figure 5. Photorespiration in 9-d-old wild-type and dct2-1 plants.
Supplemental Table 1. Absolute concentrations of maize leaf photosynthetic intermediates.
Supplemental Table 2. List of primers used for the DCT2 reverse genetic screening.
Supplemental Table 3. Number of RNA-seq raw reads.
Supplemental Data Set 1. RNA-seq data for wild-type and dct2-1 plants.
Supplemental Data Set 2. Measured mass isotopomer abundances from 13CO2 labeling experiments and peak area from 14C labeling experiments.
Acknowledgments
We acknowledge the support of the National Science Foundation (EF-1105249; IOS-092270, IOS-1127017), the Department of Energy (DE-AR0000202), and the USDA-ARS. Mass spectrometry and microscopy were performed in the Proteomics and Mass Spectrometry Core and Integrated Microscopy Core Facilities at the Donald Danforth Plant Science Center using instrumentation in part supported by National Science Foundation grants (DBI-1427621 and DBI-0521250 for acquisition of QTRAP LC-MS/MS instruments). We thank Greg Ziegler for help with the R scripts, Kaitlin McNally for help with the screen, and Kevin Ahern for help with the field genetics. Any product or trademark mentioned here does not imply a warranty, guarantee, or endorsement by the authors or their affiliations over other suitable products.
AUTHOR CONTRIBUTIONS
S.W., F.M., K.F., J.G., H.B., Y.S., M.T., D.K.A., and T.P.B. planned and performed the experiments. S.W., F.M., M.T., D.K.A., and T.P.B. analyzed the results and wrote the manuscript.
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.plantcell.org) is: Thomas P. Brutnell (tbrutnell{at}danforthcenter.org).
↵1 These authors contributed equally to this work.
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Glossary
- BS
- bundle sheath
- M
- mesophyll
- NADP-ME
- NADP-dependent malic enzyme
- NAD-ME
- NAD-dependent malix enzyme
- PEPCK
- phosphoenolpyruvate carboxykinase
- PEP
- phosphoenolpyruvate
- OAA
- oxaloacetate
- 3-PGA
- 3-phosphoglyceric acid
- TEM
- transmission electron microscopy
- 2-OG
- 2-oxoglutarate
- LC-MS/MS
- liquid chromatography-tandem mass spectrometry
- IRGA
- infrared gas analyses
- 2-PG
- 2-phosphoglycolate
- GC-MS
- gas chromatography-mass spectrometry
- CIM
- chloroplast isolation medium
- FPKM
- fragments per kilobase of transcript per million mapped reads
- Received June 5, 2015.
- Revised December 29, 2015.
- Accepted January 21, 2016.
- Published January 26, 2016.