Running Title: Protein expression atlas of BS and M maize chloroplasts

Chloroplasts in differentiated bundle sheath (BS) and mesophyll (M) cells of maize leaves are specialized to accommodate C4 photosynthesis. This study provides a reconstruction of how metabolic pathways, protein expression and homeostasis functions, are quantitatively distributed across BS and M chloroplasts. This yielded new insights in cellular specialization. The experimental analysis was based on high accuracy mass spectrometry, protein quantification by spectral counting and the first maize genome assembly. A bioinformatics workflow was developed to deal with gene models, protein families and gene duplications related to the polyploidy of maize; this avoided over-identification of proteins and resulted in more accurate protein quantification. 1105 proteins were assigned as potential chloroplast proteins, annotated for function, and quantified. Near complete coverage of primary carbon metabolism, starch and tetrapyrole metabolism, as well as excellent coverage for fatty acid synthesis, isoprenoid, sulfur, nitrogen and amino acid metabolism, was obtained. This showed e.g. quantitative and qualitative cell-type specific specialization in starch biosynthesis, Arg synthesis, N-assimilation, initial steps in S-assimilation. An extensive, overview of BS and M chloroplast protein expression and homeostasis machineries (>200 proteins) demonstrated qualitative and quantitative differences between M and BS chloroplasts, and BS-enhanced levels of the specialized chaperones ClpB3 and HSP90 that suggest active remodeling of the BS proteome. The reconstructed pathways are presented as detailed flow diagrams including annotation, relative protein abundances and cellspecific expression pattern. Protein annotation and identification data, and projection of matched peptides on the protein models, are available online through the Plant Proteome Database, PPDB. INTRODUCTION Plants can be classified as C3 or C4 species based on the primary product of carbon fixation in photosynthesis. The primary product of carbon fixation in C4 plants is a four-carbon compound (oxaloacetate; OAA), but a three-carbon compound (3-phosphoglycerate; 3PGA) in C3 plants. In leaves of C4 grasses such as maize (Zea mays), photosynthetic activities are partitioned between two antomically and biochemically distinct bundle sheath (BS) and mesophyll (M) cells. A single ring of BS cells surrounds the vascular bundle, followed by a concentric ring of specialized M cells, creating the classical Kranz anatomy. Active carbon transport (in the form of C4 organic acids) from M cell to BS cells and specific expression of Rubisco in the BS cells, allows Rubisco, the carboxylating enzyme in the Calvin cycle, to 3 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. operate in a high CO2 concentration. The high CO2 concentration suppresses the oxygenation reaction by Rubisco (and the subsequent energy wasteful photorespiratory pathway), resulting in increased photosynthetic yield, and more efficient use of water and nitrogen. The history of C4 research is described in (Nelson and Langdale, 1992; Sage, 1999; Edwards et al., 2001). Currently, there is renewed interest in C4 photosynthesis, stimulated in part by the potential use of C4 plants as a source of biofuels (Carpita and McCann, 2008) and the genetic engineering of C4 rice (Sheehy, 2007; Hibberd et al., 2008; Taniguchi et al., 2008). The use of new genomics and/or proteomics tools have resulted in new insights into cellular differentiation in C4 plants (Majeran, 2009). Proteins are responsible for most cellular functions and knowing their abundance, celltype specific expression patterns, and subcellular localization is essential to understand C4 differentiation. Previously, we published a quantitative analysis of purified M and BS chloroplast (soluble) stromal proteomes in which BS:M protein accumulati on ratios for 125 accessions were determined; this covered a limited range of plastid functions, although it enabled the integration of information from previous studies (Majeran et al., 2005). A subsequent complementary quantitative proteomics study, using nanoLC-LTQ-Orbitrap mass spectrometry and label-free spectral counting complemented with other techniques, identified proteins in BS and M thylakoid and envelope membranes of maize chloroplasts, and determined cell-type specific differences in: (i) the protein assembly state and composition of the four photosynthetic complexes and of a new type of NADPH dehydrogenase (NDH) complex; (ii) the auxiliary functions of the thylakoid proteome; and (iii) protein and metabolite transport functions of M and BS chloroplast envelopes (Majeran et al., 2008). Comparative mass spectrometry analysis of chloroplast envelope membranes from leaves of pea (Pisum sativum), a C3 species, and from M chloroplasts of maize, showed an enrichment of several known and putative translocators in the maize M envelopes (Brautigam et al., 2008). The conclusions of these proteome analyses are summarized in (Majeran, 2009). Whereas these proteomics studies provide significant progress in understanding the organization of C4 metabolism in maize, three aspects have not been adequately addressed: i) the stromal proteomes of BS and M chloroplasts likely contain each more than 1500 proteins but the BS:M ratios for only ~125 proteins were quantified, resulting in very limited coverage of several important secondary metabolic pathways such as sulfur, fatty acid, amino acid and nucleotide 4 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. metabolism, ii) information about relative concentrations of stromal proteins in BS and M chloroplasts is lacking, but is needed as a basis for quantitative modeling and metabolic engineering of C4 photosynthesis and other metabolic pathways; the growing ‘toolbox’ of proteomics and mass spectrometry now allows for such quantitative analyses (Bantscheff et al., 2007; Kumar and Mann, 2009), iii) the soluble (Majeran et al., 2005) and membrane (Majeran et al., 2008) proteome data sets were analyzed by different techniques and mass spectrometers, mostly due to the improvement of commercial mass spectrometers in that time frame. Therefore, it is difficult to understand the quantitative relationships between these datasets. The current study addresses these three aspects. So far, maize proteome analyses used essentially ZmGI maize assemblies (the Zea mays Gene Index) based on ESTs, combined with a limited amount of additional DNA sequence information. The ZmGI was originally generated by TIGR, and subsequently supported by the computational Biology and Functional Genomics Laboratory (http://compbio.dfci.harvard.edu/index.html). This ZMGI database did not have annotated gene models (for proteome analysis, the DNA sequences were searched in all six reading frames) and low expressed genes were likely underrepresented. In our most recent BS-M chloroplast analyses (Majeran et al., 2008), as well as a maize envelope analysis (Brautigam et al., 2008), the mass spectrometry data were searched against ZmGI v16.0 or v17.0. Since that time the maize genome has been sequenced (using a ‘BAC’ approach), a physical map was created (the maize accessioned golden path AGPv1), and its first assembly with gene coordinates and predicted proteins was just very recently released (June 2009; http://ftp.maizesequence.org/release4a.53/sequences/) and published (Schnable et al., 2009). This release contains 32,540 genes with 53,764 gene models; most of the gene models are evidence-based. The new maize genome assembly is expected to improve maize proteome analysis with more accurate protein identification and quantitative assessment of protein expression patterns. This also allows determination of N-terminal localization signals, which was rarely possible from EST assemblies as N-termini were often lacking. This study presents a quantitative protein expression atlas of differentiated maize leaf M and BS chloroplasts using high resolution and mass accuracy mass spectrometry (using a LTQOrbitrap) and the new maize genome assembly. Three biological replicates of stromal proteomes of isolated BS and M chloroplasts were analyzed. Quantification was carried out based on the 5 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. ‘spectral counting’ method (Zybailov et al., 2005; Bantscheff et al., 2007; Choi et al., 2008; Zybailov et al., 2008), using a sophisticated bioinformatics ‘workflow’ in particular to deal with gene duplications and extended gene families observed in polyploids such as maize. These new stromal datasets were combined with a re-analysis of our recent BS and M membrane proteome datasets (Majeran et al., 2008) against genome 4a53. Compared to previous maize leaf proteome analyses, the current study provides an integrated overview of both primary and especially secondary metabolism, as well as chloroplast gene expression and protein biogenesis, in far greater depth. The reconstructed pathways will be presented as figures that include quantitative protein information; pathways include: primary carbon metabolism, starch metabolism, nucleotide metabolism, fatty acid and lipid biosynthesis, chlorophyll, heme and carotenoid synthesis, N-assimilation. We briefly comment on the use of the new maize genome assembly for proteome analysis. All matched peptides are projected on the predicted protein models via the Plant Proteomics Database (PPDB at http://ppdb.tc.cornell.edu/). Interactive functional annotation, chloroplast localization assignments, as well as details of protein identification are also available via PPDB. RESULTS AND DISCUSSION Purified BS and M chloroplast stroma, analyzed by 1DE-SDS PAGE and nanoLC-Orbitrap mass spectrometry M and BS chloroplasts were purified from the tip of the 3 leaf of 12-14 day old maize plants in which the 4 leaf was emerging, as in (Majeran et al., 2005). The soluble stromal proteomes were collected, selected for purity and analyzed by 1D-SDS-PAGE, followed by in-gel digestion and analysis of extracted peptides by nanoLC-Orbitrap mass spectrometry. The complete analysis was carried out in three independent biological replicates. In total 144 MS runs were carried out and all spectral data were searched against maize genome assembly release 4a53, supplemented with the mitochondrial and chloroplast genomes. We identified 1662 protein models corresponding to 954 protein accessions when counting only one model per gene (see further below) (Suppl. Table 1). 6 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Purified BS and M chloroplast membranes, analyzed by BNPAGE and nanoLC-Orbitrap mass spectrometry Previously, we determined the BS and M chloroplast membrane proteomes using protein separation by native gels, followed by tryptic digestion and extensive MS/MS analysis by LTQOrbitrap (Majeran et al., 2008). Since no genome assembly was available at that time, we searched the data against the ZmGI assembly (v16.0). Here, we researched these MS data against the maize genome release 4a53. We identified 1219 protein models corresponding to 882 protein accessions when counting only one model per gene (see further below) (Suppl. Table 1). Integration of stromal and membrane datasets and selection of the best gene model The workflow of the proteome analysis is summarized in figure 2. Combining the stromal and membrane datasets identified 2439 maize gene models and 1429 protein accessions when counting only one model per gene (Suppl. Table 1). In the new maize genome annotation, many genes have more than 1 gene model. In such cases, we selected the protein form (gene model) that had the highest number of matched spectra (SPC) across all experiments; if two gene models had the same number of matched spectra, the model with the lowest digit was selected. Within each protein report page in PPDB, ‘pop-up windows’ display the gene models for each protein with the peptide count projected on the exons and details for matched peptides projected on the primary amino acid sequence. This will allow the user to determine the significance of the different gene models. BLAST search of the 1429 maize proteins against the Arabidopsis proteome and rice genome resulted in 1017 Arabidopsis and 1079 rice homologues; 47 or 48 are chloroplast-encoded in Arabidopsis and rice, respectively. Relative amounts and concentrations of proteins in BS or M cells The relative amount (mass) of each identified protein within each replicate was calculated based on the adjusted number of matched MS/MS spectra (adjSPC), normalized by the sum of adjusted SPC in the replicate, yielding nadjSPC. AdjSPC are the sum of unique SPC and a proportional distribution of shared SPC, using the ratio of unique SPC to determine this distribution; we previously developed and tested this strategy for Arabidopsis proteomes (Zybailov et al., 2008). The relative concentration for each identified protein was calculated as the normalized Spectral 7 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Abundance Factor (nSAF), which was calculated from adjSPC weighted for the number of theoretical tryptic peptides with a relevant length (‘observable peptides’) (Zybailov et al., 2006). To better understand the distribution and expression range of the identified protein population, we determined their frequency distribution of the relative concentration using Log10(nSAF) values and a bin size of 0.50 (Fig. 1B). Bins in the range of -1.5 to -3 with the ~110 most abundant proteins, were strongly dominated by proteins involved in photosynthesis and the C4 shuttle. Bins in the range of -5.5 to -7, with the ~205 proteins of lowest abundance, contained mostly proteins with less than 5 matched spectra, frequently with most of the spectra shared to other proteins; thus this group of low abundant proteins is enriched in low expressed members of groups of homologues, possibly indicating that these are pseudogenes, as well as false positive identifications (see below). Removal of low scoring proteins Our workflow has a strictly controlled false positive peptide identification rate and we use high resolution (100,000), high mass accuracy (6 ppm) MS data; this workflow helps to avoid false positive identifications. However, the high complexity of the maize genome and presence of repetitive DNA and the fact that we are using the first draft genome sequence, could lead to identifications of less meaningful protein accessions (eg missassemblies, pseudogenes, etc). Therefore for further analysis of BS and M chloroplast functions, we removed those protein accessions with only one matched MS/MS spectrum (in total 100 accessions). In addition we removed those protein accessions that were identified with only one amino acid sequence (irrespective of charge state and possible post-translational modifications) if the sequence had less than 10 amino acid residues and contained isoleucine (I) or leucine (L); these two residues have the same mass (they are isobaric) and can thus not be distinguished by MS. These steps removed 149 proteins, mostly representing pseudogenes, false positive identifications and proteins expressed at very low levels; they represented 0.07% of the calculated protein mass (% of total nadSPC) (see Suppl. Table 2). Assignment of chloroplast localization Since we were interested in BS and M chloroplast differentiation and function, we evaluated the remaining identified proteins for chloroplast localization. For assignment of a protein to the 8 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. chloroplast, we used a combination of mass spectrometry–based scores (chloroplast proteins should generally have higher scores in chloroplast-enriched fractions than in total leaf fractions), and known localization of the predicted best homologue in Arabidopsis or rice, in addition to the known or predicted maize protein function. We did pay careful attention to localization assignment for proteins that are members of groups of closely related identified homologues. We assigned 974 proteins to the chloroplast, 175 proteins were assigned to other subcellular locations and were considered contaminants, and for the remaining proteins we could not assign a subcellular location (Suppl. Table 2). These three categories had respectively 99%, 0.6%, 0.3% of the total nadjSPC, indicating that assigned contaminations made up less than 0.6% of the total protein mass. The actual contamination was lower since highly expressed (chloroplast) proteins are underestimated in their abundance using the spectral counting technique (Zybailov et al., 2008). We then analyzed the frequency distribution of relative protein concentration (based on Log10(nSAF)) for these three groups of proteins. This showed that the chloroplast proteins spanned 4 orders of magnitude, peaking at -3.5 to -4, whereas the contaminant proteins spanned about 3.5 orders of magnitude in abundance, peaking at -5.5 to -6 (Fig. 1B). For further analysis, we removed the contaminant proteins. The presence of organellar genes in the nuclear genome assembly In addition to the nuclear genome, plastid and mitochondria also have a genome and we merged these sequences with the 4a53 nuclear genes when we searched the mass spectral data. Indeed, we identified 47 chloroplast-encoded proteins and 1 mitochondrial-encoded protein (Suppl. Table 2). However, we noted that several of these chloroplast-encoded proteins had many shared MS/MS spectra matching to maize 4a53 (nuclear genome) accessions. BLAST searching of these maize accessions against Arabidopsis and rice, showed they mostly matched to chloroplastencoded Arabidopsis and rice proteins. These 4a53 genes could represent pseudogenes from fragments of plastid genome inserted into the nuclear chromosomes, or could result from DNA sequencing of contaminating chloroplasts DNA. Examples are the observation of seven maize 4a53 genome accessions for the chloroplast-encoded Rubisco large subunit (RBCL) in addition to the chloroplast accession (NP_043033), and four maize 4a53 genome accessions for the chloroplast-encoded PSII subunit cytb559α in addition to the chloroplast accession (NP_043041). For quantification purposes we grouped these redundant accessions as will be described further 9 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. below (Section: Grouping of identified protein accessions to deal with gene duplications and extended gene families in maize for quantification). Functional annotation of proteins, differentiating between C3 and C4 enzymes and splicing The remaining proteins were assigned a function using the MapMan bin classification system (Thimm et al., 2004), similar to its use in previous maize studies (Majeran et al., 2005; Majeran et al., 2008). Moreover, the MapMan classification system is well integrated in our PPDB for both Arabidopsis and maize proteome data (Sun et al., 2009). We added a new bin (assigned 1.5 PS.C4 malate shuttle) for proteins involved in the C4 shuttle (e.g. phosphoenolpyruvate carboxylase PepC, pyruvate phosphate dikinase PPDK, malate dehydrogenase MDH, PPDK regulatory protein PPDK-RP); importantly, based on quantitative BS-M expression patterns, we were also able to differentiate between proteins specialized in the C4 functions as compared to non-C4 functions (e.g. PPDK and MDH). For instance, we identified 2 PPDK accessions, GRMZM2G011507 (PPDK-C4) and GRMZM2G097457 (PPDK-C3). PPDK-C4 is extremely abundant, with a BS/M ratio of 0.56, whereas less abundant PPDK-C3 is nearly exclusively localized in BS cells (BS/M ratio is 9.7). Interestingly, we identified only one maize gene accession for PPDK regulatory protein (PPDK-RP) GRMZM2G004880. PPDK-RP is enriched in M chloroplasts (BS/M =0.28), indicating its specificity towards the regulation of the mesophyll-localized C4-type PPDK. There are no obvious homologues in the 4a53 genome and indeed only one maize PPDK-RP was identified by bioinformatics analysis and cloning (Burnell and Chastain, 2006). In Arabidopsis there are two genes for PPDK-RP (RP1 and RP2, respectively At4g21210 and At3g01200). RP1 is similar to the maize C4 type PPDK-RP and is plastid localized, whereas RP2 has no detectible activity and is localized to the cytosol (Chastain et al., 2008). For more details on C4 and non-C4 NADP-MDH enzymes, see (Maurino et al., 2001; Tausta et al., 2002). The identified proteome provides high coverage of secondary metabolism, plastid gene expression and chloroplast protein homeostasis Figure 1C shows the distribution of the number of proteins (total 1105 proteins) or protein mass involved in i) primary carbon metabolism and photosynthesis, ii) secondary metabolism, iii) miscellaneous and unknown functions, iv) plastid gene expression and protein homeostasis and 10 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. v) membrane transport. Each of the first four groups involved 23-25% of the identified proteins, whereas about 4% of identified proteins were involved in transport of metabolites across the chloroplast envelopes (Fig 1C – upper panel). In terms of protein biomass the majority (~66%) was invested in primary metabolism, with equal distribution across secondary metabolism and plastid protein homeostasis (each 13%) (Fig 1C lower panel). A more detailed breakdown of chloroplast functions is shown in figure 1D and shows high coverage of functions not much observed in maize chloroplasts in previous studies. In particular, the protein synthesis components involved in expression of chloroplast-encoded proteins were very well covered – this included most of the ribosomal proteins, many of the tRNA synthetases, as well as initiation and elongation factors (Fig. 1D). Even more rewarding was the identification of 132 proteins involved in various aspects of post-translational chloroplast protein homeostasis, including about 50 processing peptidases, amino-peptidases, proteases, as well as soluble methione sulfoxide reductases involved in protein repair, peptide deformylase, as well as several phosphatases and kinases. Most if not all members of the Clp protease system in maize were identified; while they are now well characterized in Arabidopsis chloroplasts (Sjogren et al., 2006; Kim et al., 2009; Zybailov et al., 2009), they appeared elusive in maize. Excellent coverage was obtained for redox regulators and ROS detoxification enzymes (e.g. peroxiredoxins), nitrogen, sulfur and amino acid metabolism and other secondary metabolic pathways, including fatty acid synthesis, isoprenoid, tetrapyrole, nucleotide metabolism (Fig. 1D). Whereas only a very small percentage of protein mass was invested in isoprenoid synthesis and derived products, as well as cofactor and vitamin biosynthesis (less than 1%), we obtained a good coverage of several of these pathways; for instance most enzymes of the methylerythritol phosphate pathway (MEP) pathway and multiple enzymes in thiamine (vitamin b1) and riboflavin (vitamin b2) synthesis were identified. Thus the combined analysis of membrane and soluble fractions of isolated BS and M chloroplasts provided a good coverage of many of the chloroplast functions. Therefore, the quantitative comparison of the BS and M profiles should thus provide meaningful and new insights into C4 driven differentiation and fulfill our objective to obtain an integrated, quantitative overview of the differentiated state of BS and M chloroplast functions in the maize leaf. The differentiated functional state of BS and M chloroplasts 11 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Table 1 compares protein mass investment (based on nadjSPC) of BS and M chloroplasts into the different chloroplast functions (Table 1). About 39% (M) to 38% (BS) of the total chloroplast proteome was invested in the photosynthetic apparatus and cyclic electron flow located in the thylakoid membrane (Table 1). Between 23% (M) and 31% (BS) was invested in the Calvin cycle, the C4 shuttle and carbonic anhydrases. About 11% (M) and 10% (BS) was invested in plastid gene expression, protein folding, processing and proteolysis. About 5% (M) and 3% (BS) was invested in redox regulation. 1.9% (M) to 1.6% (BS) was invested in transport functions. Some 6% (M) and 5% (BS) was invested in proteins for which the function is unknown. About 14% (M) and 11% (BS) was invested in all remaining functions (Table 1). To better appreciate the cell-type specific differences in molecular functions, we further broke down these protein investments for the membrane and soluble fractions (Table 1) – for a detailed description, that also highlights the most abundant proteins for the various functions, we refer to Suppl. Text. . Grouping of identified protein accessions to deal with gene duplications and extended gene families in maize for quantification Maize is a polyploid with a far larger genome size (2800 Mbp) than Arabidopsis (130 Mbp) and rice (430 Mbp), and maize has a significantly larger amount of repetitive sequences (Schnable et al., 2009). The complexity of the maize genome and its predicted proteome require an extra effort for maize proteomics, in particular to deal with closely related identified proteins. These closely related proteins could represent true duplicated genes, pseudogenes or could be artifacts of the genome assembly. To deal with this complexity of the maize genome for quantification of BS and M protein expression, we grouped proteins that shared more than ~80% of their matched SPC. Grouping was done by generation of a similarity matrix through calculation of the dice coefficient between each pair of identified proteins based on matched SPC, followed by clustering of the proteins using MCL software (Enright et al., 2002), followed by manual evaluation (see Material and Methods). In total 313 proteins were placed in 131 groups (Fig. 2). This grouping avoids overinterpretation of differences observed between M and BS chloroplast proteomes; this strategy was also very useful for Arabidopsis (Rutschow et al., 2008)(Zybailov et al., 2009). The ‘relationships’ between proteins that shared matched MS/MS spectra are visible through PPDB. Quantification of proteins with a low number of adjSPC within a replicate (i.e. below ~10 12 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. adjSPC) is generally less accurate (Kim et al., 2009; Zybailov et al., 2009). 378 proteins had less than 40 adjSPC, 179 had between 40 and 100 adjSPC and 366 had more than 100 adjSPC (Fig. 2), and we generally consider these three groups as low, medium and high confidence quantifications, respectively (Suppl. Table 3). In the remaining sections, metabolic pathways are reconstructed, and the distribution of specific proteins in the various processes and metabolic pathways over the BS and M chloroplasts is discussed. We will first discuss the chloroplast gene expression and protein homeostasis machineries to understand assembly and maintenance of the differentiated chloroplast proteomes. Furthermore we focus on those metabolic pathways that were not (or poorly) covered in previous maize studies, in particular starch metabolism, nucleotide metabolism, sulfur and nitrogen assimilation, isoprenoid and tetrapyrole metabolism. We will also analyze the distribution of relative concentrations of the proteins, as this will be valuable for future modeling of metabolic fluxes. In these various plots and analyses we will use nSAF values, as this is the best approximation of relative protein concentration, as well as BS/M accumulation ratios. The data will be presented as figures for plastid gene expression, protein synthesis and homeostasis (Fig. 3); thylakoid light reactions (Suppl. Fig. 1), envelope transporters (Fig. 4), primary carbon metabolism (Fig. 5), starch metabolism (Fig. 6), nucleotide metabolism (Fig. 7), fatty acid and lipid biosynthesis (Fig. 8), chlorophyll, heme and carotenoid synthesis (Fig. 9), Nassimilation (Fig. 10), S-assimilation (Fig. 11), and tables for redox regulation and ROS defense (Table 2) and for amino acid metabolism (Table 3). More details regarding e.g. accession numbers, groupings of accessions, Mapman bin numbers, nSAF values, matched MS/MS spectra and more, are available in Suppl. Table 3. Plastid gene expression, protein synthesis and homeostasis The contribution of plastid gene expression and protein homeostasis on control of BS and M differentiation is entirely unclear. For instance, levels of chloroplast-encoded (and nuclearencoded) PSII subunits are lower in BS than in M chloroplasts, whereas chloroplast-encoded NDH subunits are higher in BS chloroplasts. However, it is not clear if levels of these chloroplast-encoded proteins are regulated through control of transcription (or even chromosome copy number), mRNA processing and stability, translation, or post-translationally, through proteolysis. Furthermore, levels of imported nuclear-encoded proteins within the BS or M 13 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. chloroplast could be controlled through proteolysis. Both chloroplast and nuclear-encoded proteins share intra-plastid protein sorting, folding and assembly machineries. After careful evaluation for function and location, we quantified 221 proteins (and protein groups) involved in chloroplast biogenesis and protein homeostasis, assembled in five categories, namely protein synthesis (77 proteins), proteolysis and processing (42 proteins), import and post-translational modifications (34 proteins), folding (38 proteins) and RNA/DNA interaction (31 proteins). Figure 3 displays the BS/M ratio (based on nSAF), the number of matched adjSPC, and relative abundance of proteins (as a color scale) in these various functional classes. 114 proteins were only found in the stroma, whereas 72 were found in both membrane and stromal fractions, but with a wide range of distribution (i.e. between 3000-fold enriched in the membrane fraction and 100-fold enriched in the stromal fraction). The systematic quantitative comparison of BS and M chloroplast proteins in these processes did identify several strongly differentially expressed proteins (marked with an asterisk in Fig. 3); these may contribute to BS and M specialization. We will highlight selected candidate proteins, with emphasis on soluble proteins and comment on functional implications and follow-up analyses (see Suppl. Table 3 for details). Interactors with the plastid chromosome There is very little understanding as to how plastid chromosome copy number and organization influence transcription and contribute to BSM chloroplast differentiation. Therefore, we were pleased to identify eight DNA interacting proteins (pTAC or nucleoid proteins) and two DNA repair enzymes. The abundance levels of these proteins spanned more than 2 orders of magnitude (based on nSAF), with two membranebound nucleoid associated proteins, pTAC16 and MFP1 being by far the most abundant, with respectively 1679 and 656 matched MS/MS spectra. pTAC16 of unknown function was two-fold enriched in the mesophyll, whereas MFP1 was 2-fold enriched in the BS chloroplasts. MFP1 is a coiled-coil DNA-binding protein and is believed to play a role in anchoring the plastid DNA to the envelope and thylakoid membrane; its expression is tightly correlated with the accumulation of thylakoid membranes (Jeong et al., 2003). Two soluble putative DNA repair enzymes, auvrB/uvrC motif-containing protein and a deoxyribodipyrimidine photolyase follow in abundance and have BS/M ratios of respectively 0.51 and 0.99. The next three abundant proteins were homologues of TCP34, pTAC5 and pTAC17; TCP34 and pTAC5 homologues were only found in the membrane fractions and pTAC17 mostly in the soluble phase. The Arabidopsis 14 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. homologues of these pTAC proteins were all identified in a highly enriched plastid chromosome preparation, but their precise function is not clear (Pfalz et al., 2006; Weber et al., 2006). Targeted nucleoid and functional analyses will be needed to determine the contribution of transcriptional regulation to chloroplast BS and M differentiation. Regulation of transcription and RNA metabolism We identified 21 proteins in this category, including six RRM domain proteins, and four ribonucleases in the soluble stromal fractions covering 2-3 orders of expression with the number of matched MS/MS spectra ranging from 1400 to just 3 (Fig. 3). The most abundant proteins were homologues of CP33, CP31, CP29, CSP41A and CSP41B. Homologues for several of these proteins have been characterized in Arabidopsis – examples are RRM protein CP31A (Tillich et al., 2009) and CSP41 (Beligni and Mayfield, 2008; Bollenbach et al., 2009). CSP41B-2 (BS/M =2.5) and CSP41A (BS/M 2.1) and the 3-5’ exoribonuclease RIF10 (BS/M = 4.9) and a DUF740 protein (BS/M =3.5) were all higher expressed in BS chloroplasts, whereas several other, such as two SET domain proteins and a S1 RNA biding proteins were much more expressed in M chloroplasts (BS/M = 0.2, 0.1 and 0.06). The chloroplast translational machinery. We quantified 76 proteins (and protein groups) that are part of the chloroplast translational machinery, including 18 tRNA-synthetases, two initiation (IF2, IF3) and five elongation factors (BipA, TU, G and P types), a ribosome recycling factor and a peptide chain release factor, as well as 14 subunits of the 30S ribosomal particle and 32 subunits of the 50S particle and three plastid specific ribosomal proteins (PSRP1,2,3). Relative protein concentrations spanned three orders of magnitude, with the most abundant protein being EF-TU-1 with both elongation and chaperone function (3163 matched MS/MS spectra). Chloroplast ribosome levels were about 3-fold higher in M chloroplasts than BS chloroplasts (median BS/M ratio for individual subunits was 0.32). Interestingly, the initiation and elongation factors were more equally distributed across BS and M, with the exception of the typA/BipA elongation-factor like protein. TypA/BipA EF is a specialized ribosome associated translation factor (Wang et al., 2008) suggested to be required in stress response; interestingly we observed BipA EF to be strongly induced in Arabidopsis chloroplast Clp protease mutants (Kim et al., 2009; Zybailov et al., 2009). The 18 tRNA synthetases were higher in M chloroplasts, with a median BS/M ratio of 0.14. Overall, these data suggest higher translational activity in M chloroplasts than BS chloroplasts, particularly at the thylakoid surface. 15 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. When comparing the stromal and membrane proteomes, we calculated that nearly 40% of ribosome protein mass (based on NadjSPC) was found in the membrane fractions. Interestingly, the M membranes fractions contain 20-fold more ribosomal protein than the BS membranes, which is very striking when compared to the ~2-fold difference between the M and BS stroma. Chloroplast ribosomes are known to associate with the thylakoid membranes, and indeed many chloroplast–encoded thylakoid proteins are synthesized at the membrane surface and cotranslationally inserted (Margulies and Michaels, 1975; Klein et al., 1988; van Wijk et al., 1996; Rohl and van Wijk, 2001). We suggest that the very low BS/M ratio of thylakoid-bound ribosomes reflect a much higher demand for synthesis of thylakoid proteins in M chloroplasts, most likely due to high M abundance of PSII subunits and the relative short life-time of PSII reaction center proteins due to light-induced damage. The BS-enriched NDH complex also contains many chloroplast-encoded proteins, but the relative concentration of the NDH complex, and likely also the turn-over rate, is several fold lower than the PSII complex, thus contributing much less to overall chloroplast translation. Protein processing and proteolysis We quantified 42 proteases and processing peptidases, 26 of which were quantified with medium or high confidence. These include 22 stromal, six thylakoid lumen proteins, nine integral thylakoid proteases and three proteases that we assigned to the inner envelope membrane. As most membrane proteins were discovered and discussed previously when searching the ZmGI database (Majeran et al., 2008), we focus here on the 22 soluble proteases and peptidases. 12 of the soluble protease were members of the Clp protease system, and based on a multi-align analysis with the Arabidopsis Clp family, we annotated the maize Clp proteins. The Clp protease system is the most abundant soluble protease in Arabidopsis chloroplasts (Peltier et al., 2004) and in pea etioplasts (Kanervo et al., 2008) and consists of a proteolytic tetradecameric barrel-like structured ClpPR complex to which two small ClpT subunits tightly associate (Peltier et al., 2004). ClpC1,C2 chaperones are assumed to deliver substrate to the core protease complex (Adam et al., 2006). We were surprised to find that the most abundant ClpPR subunit in maize chloroplasts was a homologue of Arabidopsis mitochondrial ClpP2. Clearly this maize ClpP2 protein is not mitochondrial and this finding warrants a more in-depth phylogenetic analysis of the Clp protease family in plants. The average and median BS/M ratio of the Clp system was respectively 0.8 and 0.7; this rather equal distribution across BS and M chloroplasts is consistent with the idea that the Clp system serves a 16 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. general housekeeping protease, unlike some of the proteases with narrow functions and extreme BS/M ratios, such as SPPA. The slight bias (25-40%) towards M accumulation may relate to specific control functions by the Clp protease system of plastid gene expression; this is certainly worth-while investigating further. Other abundant proteases were Prep1 (likely involved in degradation of cleaved cTPs), several amino peptidases (eucyl, glycyl, glutamyl, M24) and a Lon protease (LON2). Several of these soluble proteases showed a strong preferential distribution for either cell type; in particular glutamyl endopeptidase (cGEP) and abundant glycyl aminopeptidase (M1) were respectively 4fold and 2-fold higher in the BS stroma, whereas stromal DegP2, the very abundant eucyl aminopeptidase LAP1 and M24 aminopeptidase APP2 were respectively 3-fold, 4-fold and 5fold higher in M chloroplasts. The substrates of these proteases are unknown but the preferential accumulation in one cell-type suggests a narrow set of substrates. Protein sorting We identified the major soluble and membrane components that control protein import, sorting and translocation in the thylakoid (Tic110,40,55; SRP43/54,SecA/Y and TatC and HCF106). Tic110, Tic40, SecY and SecA were each identified with more than 50 spectral counts and their BS/M ratio could be determined with confidence. The envelope import components, as well as both subunits of the SRP particle and soluble SecA were clearly higher in M chloroplasts, whereas SecY was equally distributed. Since most of the SecA dependent abundant lumenal proteins are components on M-enriched PSII complex, the strong M accumulation is logic. SecY is the general thylakoid import channel for membrane proteins and its equal distribution across the two cell types is consistent with its general role. Protein assembly factors. We also identified 13 soluble and membrane proteins involved assembly of thylakoid complexes; these included several factors that function for very specific complexes (e.g. for PSII HCF136, LPA1; for PSI – PYG7, YCF4), and at least three factors involved in biogenesis of 2Fe-2S complexes (NFU1,2,3) (see also the section on S-assimilation, below). Factors for PSII specific complexes were several fold higher in M chloroplasts and those involved in PSI biogenesis were 30-40% higher in BS see further also in (Majeran et al., 2008). Thus the BS/M accumulation ratios of the PSI and PSII specific assembly factors correlate well with the BS/M ratios of PSI and PSII themselves, indicating that expression of these assembly factors must be well coordinated with demand. Strikingly, the maize homologue of Arabidopsis stromal protein HCF101 (quantified with 69 adjSPC) involved in biogenesis of 4Fe-4S clusters 17 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. (not 2Fe-2S) in PSI (Stockel and Oelmuller, 2004), as well as ferredoxin-thioredoxin reductase (FTR) (Lezhneva et al., 2004), showed 10-fold higher M accumulation. Unlike PSI, FTR is more highly expressed in M chloroplast than BS chloroplasts, and further studies on the regulation of cell-specific expression and accumulation of HCF101 may elucidate regulatory networks for chloroplast biogenesis and differentiation. Protein (un)folding and maturation We quantified 38 proteins involved in (un)folding and maturation; none had predicted transmembrane domains. These proteins included lumenal protein isomerases, general high abundant stromal chaperones (several HSP70 and their GrpE nucleotide-exchange factors and CPN60/20/10 proteins), as well as low abundant factors (eg BSD2 implicated in Rubisco assembly). As discussed previously (Majeran et al., 2008), several thylakoid lumen isomerases showed distinct preferential accumulation in BS or M thylakoids, suggesting specific adaptation or substrates. General chaperones were quite equally distributed consistent with their broad array of substrates. Clear exceptions were stromal chaperone HSP90 and ClpB3, involved respectively in protein maturation and unfolding; they were 2-6 fold higher in BS chloroplasts. Considering the much lower chloroplast translation rates in BS chloroplasts, this is surprising and potentially very important for understanding the BS-M differentiation pathways; we speculate that this relates specifically to the specialization of the BS chloroplast (see Conclusions further below). Post-translational modifiers Nuclear-encoded chloroplast proteins can be modified within the chloroplast after import, whereas chloroplast-encoded proteins can be modified during and after synthesis. We identified 2 different types of protein repair proteins, methionine sulfoxide type-A4 (MSRA4) and ribulosamine/erythrulosamine3-kinase; both were only slightly more abundant in M chloroplasts, possibly because the anti-oxidative systems within the chloroplast have sufficient capacity to prevent protein damage. We identified peptide deformylase 1A (PDF1a), involved in co-translational removal of N-terminal formyl group of methionine, with a BS/M ratio of 0.68; this higher M accumulation is again consistent with higher translation rates in M chloroplasts. The function of the very abundant and soluble methyltransferase (246 MS/MS spectra) is unknown and it has a similar BS/M ratio as PDF1a. Phosphorylation does play an important role in chloroplast metabolism and adaptation, but more systematic studies on stromal kinases and phosphatases are now just beginning to emerge (in Arabidopsis) (Schliebner et al., 2008; Reiland et al., 2009). We identified 2 protein phosphatases 18 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. 2C (PP2C), a protein tyrosine phosphatase and a putative protein kinase inhibitor. The most abundant PP2C (246 MS/MS spectra) was more than 5-fold enriched in BS chloroplasts, whereas the Zinc-binding domain protein and putative kinase inhibitor (68 MS/MS spectra) were more than 5-fold enriched in M chloroplasts. Reversible phosphorylation, in addition to redox regulation (see further below) plays an important role in regulation of plant metabolism, and the study of BS and M chloroplast specific (de)phosphorylation networks will be an important complement to our current study. The photochemical apparatus in the thylakoid membrane We obtained extensive coverage (121 proteins and protein groups) of the linear electron transport chain including PSII (33 proteins), PSI (23 proteins), the cytb6f complex (5 proteins), the ATP-synthase (9 proteins), lumenal plastocyanin (PC), five Fd proteins, three FNR1 isoforms, and six light-stress proteins with chlorophyll binding domains of the Ohp and Lil families (Suppl. Fig. 1). We also identified cyclic electron flow components of the NDH complex and NDH specific biogenesis factors (23 proteins), NDH-independent cyclic electron flow components of the PGR complex (3 PGRL1 and 2 PGR5 homologues), as well as alternative thylakoid terminal oxidases (PTOX or IMMUTANS) (2 homologues) and PIFI. We note that homologues of the new NDH subunits that we discovered previously in maize (Majeran et al., 2008), have now also been identified in Arabidopsis chloroplasts (Peng et al., 2009). Finally, we identified two state transition kinases (Stn7 and Stn8), and the thylakoid phosphoprotein TSP9 and the Ca phosphoprotein (marked in Suppl. Fig 1. under DE-P). Most of these proteins were also identified based on our previous search against the ZmGi EST assembly database (Majeran et al., 2008), and we will therefore not discuss these proteins any further. BS/M ratios for these proteins per complex or function are shown in Suppl. figure 1. Envelope transporters with known and unknown functions From our recent BS-M membrane study (Majeran et al., 2008) and general literature, combined with functional suggestions from the study of Weber and colleagues (Brautigam et al., 2008), we (tentatively) assigned functions for 26 chloroplast envelope transporters (Majeran, 2009); these included mesophyll enriched transporters MEP1,2,3,4 with unknown substrates, members of the DIT family (Dit1, Dit2 and OMT1), phosphate/triosephosphate translocators (TPT, PPT), the 19 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. maltose exporter MEX1, ATP/ADP translocator AATP1 or NTT1, and anion transporter ANTR2. A number of envelope transporters (e.g. various porins, ABC transporters) could not be assigned to specific functions. Researching our membrane data against the maize genome identified 46 transporters (52 proteins in 46 groups). We summarize thes information for these transporters with (putative) substrates in figure 4. Log2 BS/M ratios are displayed in small bar diagrams, with the bars shown in black, dark grey and light grey, which indicate proteins identified with respectively high (>100), medium (40<>100) and low number of adjSPC (<40). Proteins that are quantified with at least 40 adjSPC and enriched more than 1.5-fold in the M chloroplasts are marked in blue, whereas enzymes enriched more than 1.5 fold in the BS chloroplasts are marked in red. Relative protein abundance (based on nSAF) across both cell types are shown as colored squares. The transporter with the highest relative abundance was MEP1-1 (equally distributed between BS and M), followed by MEP3/4 (M-enriched), PPT (strong M-enriched), TPT (M enriched). Some of these transporters are also integrated with information of specific pathways, in particular with primary carbon metabolism and the C4-malate shuttle (Fig. 5), starch metabolism (Fig. 6), nucleotide and nitrogen metabolism (Figs. 7, 9). These assignments are based on previous literature and in the case of the Calvin cycle intermediates (3PGA and DHAP) and Malate shuttle substrates (PEP, OAA), these assignments are speculative; Identification of substrates for many of these transporters should be of highest priority. Calvin cycle, the C4 malate shuttle, the oxidative pentose phosphate pathway and glycolysis Whereas our previous stromal analysis, employing 2DE gels and Q-TOF based mass spectrometry analysis, identified several of the enzymes involved in the Calvin cycle and the the C4 malate shuttle (Majeran et al., 2005), the new stromal analysis using spectral counting, the LTQ-Orbitrap and the new maize genome, gives a near complete overview. Figure 5 shows an integrated overview with BS/M ratio of the Calvin cycle, the C4 shuttle (PPDK, PPDK-RP, MDH, ME), as well as the (irreversible) oxidative pentose phosphate pathway (G6PDH, Lact, 6PGDH) and the reversible pentose phosphate pathway (TKL, RPE, RPI, TA), and enzymes leading to starch biosynthesis (PGM1, PGM2, Glc6PI). Furthermore, the figure shows C3-forms of abundant C4-shuttle enzymes (NADP-MDH and PPDK). The inset above the diagram shows the relative abundance of proteins for both cell types combined. We did not find evidence for 20 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. accumulation of the specific chloroplast glycolytic enzymes (PyrK, ENI, PGlyM), nor did we find evidence for PPI-PFK or ATP-PFK involved in conversion of F6P to F16BP. We did identify PGP, one of the two chloroplast-localized enzyme involved in photorespiration, only in the BS chloroplast. Chloroplast glycerate kinase, involved in conversion of glycerate imported from peroxisomes into 3PGA is likely GRMZM2G054663, but we never identified it. In agreement with our initial analysis in 2005 (Majeran et al., 2005), our data strongly suggest that the reductive phase of the Calvin cycle, represented by GAPDHB and TPI, is enriched in the M chloroplast. The third enzyme in the reductive phase, PGK1, was also slightly higher in the M chloroplast, in contrast to the homologue PGK2, which was much higher in the BS chloroplast. Moreover, two of the reversible pentose phosphate pathway enzymes (RPI, trans aldolase (TA) -marked with an asterisk) are also (somewhat) higher in M chloroplasts. The three enzymes of the oxidative PPP (G6PDH, Lact., and 6PGDH) are more expressed in the M chloroplasts. A second low abundant isoform of Lact was higher in BS chloroplasts, but the significance is unclear, since it was only quantified by 8 MS/MS spectra. The increased levels of the OPPP enzymes in M chloroplasts suggest that carbohydrates imported from the BS cell feed the OPPP in M chloroplasts likely as a source of carbon intermediates for various pathways. Alternatively, the OPPP pathway may also be higher to provide precursors to the shikimate pathway; indeed as discussed further below (and Table 3), most enzymes in the shikimate pathway were more abundant in the M chloroplast than BS chloroplast. The preferential BS accumulation of PGM1, PGM2 and Glc6PI is not a reflection of increased rates of glycolysis, but instead reflects higher rates of starch synthesis in BS chloroplasts (see next section). The C3-type PPDK is strongly enriched in BS chloroplasts, whereas C3-type NADP-MDH is equally distributed across both cell types. Pathways for starch metabolism show strong quantitative and qualitative differences between BS and M chloroplasts Expression and distribution of starch metabolic enzymes in C4 leaves has not been systematically studied. We identified and quantified the relative abundances of 21 chloroplastlocalized enzymes involved in starch synthesis and degradation. These 21 proteins were assigned names, functions and positions in the starch metabolic pathway, based on BLAST alignments and extensive literature analysis, e.g. (Smith et al., 2005; Zeeman et al., 2007) and (Fulton et al., 21 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. 2008) (Fig. 6). We used the nomenclature developed for Arabidopsis (Smith et al., 2004). For functional interpretation, we connected the starch pathway to enzymes (PGM1,2;Glc6PI) involved in conversion between glucose-1P and glucose-6P (Glc1P;Glc6P) and fructoses (F16BP;F6P), as well as the maltose and glucose transporters (MEX1, GlcT1) (Fig. 7). BS/M protein ratios are indicated as bar diagrams and the total protein abundance (based on nSAF) across both cell types are shown as colored squares. The majority of enzymes are more abundant in BS chloroplasts than M chloroplasts and the total investment of enzymes in starch metabolism is nearly three-fold higher in BS than in M chloroplasts (see Table 1); this is consistent with several previous scattered observations (Spilatro and Preiss, 1987; Lunn and Furbank, 1997; Majeran et al., 2005; Majeran et al., 2008), and the much higher presence of starch particles in BS chloroplasts than M chloroplasts. ADP-glucose-pyrophosphorylase (AGPase), a heterotetramer of large and small subunits, is the first committed step in starch biosynthesis and is controlled by a combination of allosteric control by 3PGA and Pi, redox regulation and trehalose (Kolbe et al., 2005; Zeeman et al., 2007). We indentified two isoforms of the large subunit of AGPase (AGPL1,2) and one isoform for the small subunit (AGPS). APGL1 is nearly 10-fold more abundant than APGL2; APGL1 is about 2fold higher in BS than in M chloroplast, whereas APGL2 is likely a specific isoform adapted to M chloroplast conditions since it is >10-fold higher in M than in BS chloroplasts. APGS was ~2fold higher in BS than in M chloroplasts and likely serves both large isoforms. The product of AGPase, ADP-glucose (ADPGlc) is used by a family of starch synthases (SS) to generate linear glucose polymers named amylose, and branched glucose polymers named amylopectin. We identified four starch synthase proteins (SS), namely granule bound SS (GSS), which is needed for synthesis of long linear chains of amylose, and SSI, SSIIa and SSIIIb which generate the linear chains of amylopectin. GSS was 4.4 fold higher in BS than in M chloroplasts, consistent with a higher production of starch in BS and the requirement of GSS for amylose synthesis. It is thought that SSI is needed upstream of SSII and that SSII operates upstream of SSIII; SSI is primarily responsible for synthesis of short chains, and SSII and SSIII lengthen these chains further. SSI was equally distributed between BS and M chloroplasts, whereas the more abundant SSIIa was more than 2-fold higher in BS chloroplasts. The low abundance SSIIIb was 1.4 fold higher in BS chloroplast. Together, this strongly suggests that starch synthesized in M chloroplast has much shorter amylase chains as compared to B chloroplasts. 22 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. We identified three starch (de)branching enzymes (BEIIb1,2 and ISA2) involved in synthesis of branched glycans. BEIIb introduces α-1,6 branch points and is a so-called class II branching enzyme. Absence of class II BE, results in production of only long chain glucans and no amylopectin. ISA2 is a type of debranching enzyme (a DBE) involved in synthesis (rather than degradation) and it has been suggested that ISA2 has more a regulatory function (see for discussion (Zeeman et al., 2007)). BEIIb2 was about 100 times more abundant than BEIIb1 and ISA2. BEIIb2 was >6 fold enriched in BS chloroplasts, whereas EBIIb1 and ISA2 were not detected in M chloroplasts. Together this suggests that the starch produced in BS chloroplasts is more highly branched than starch in M chloroplasts. We identified two kinases (PWD and GWD) that catalyze the phoshorylation of a glucosyl residue of amylopectin; these two enzymes stimulate starch degradation, possibly by loosening the surface of the granule structure and/or increasing the solubility of starch (Smith et al., 2005; Zeeman et al., 2007). Both have a similar relative concentration and both were enriched (3.5-5 fold) in BS chloroplasts; these BS-enriched levels are indicative and/or consistent with the reduced accumulation of amylopectin in M chloroplasts (see further below). We also identified the maize homologue of Arabidopsis phosphoglucan phosphatase DSP4 (or SEX4) (Kotting et al., 2009) and DSP4 was 3-fold enriched in M chloroplasts. DSP4 phosphorylation activity is needed to allow effective cleavage hydrolysis of glucans by BAM3 and ISA3. The preferential M accumulation of DSP4 is puzzling and suggests a specific adaptation of starch degradation. There are several starch breakdown pathways in leaves, and they result in maltose, glucose or Gcl1P (Zeeman et al., 2007). We were able to assign maize homologues to each of these pathways in BS chloroplasts. In C3 leaves such as Arabidopsis, the maltose is the main breakdown product and export product of transient starch in leaves during the dark period. Based on the relatively high levels of ISA3 as compared to DPE1 (~15 fold higher), it is likely that BS chloroplasts also use maltose as main export product in the night. However, our observations of high levels of DPE1 (comparable to ISA3 in BS chloroplasts) and its low BS/M ratio (Fig. 6), suggests that in M chloroplasts the dominant end product for starch degradation is glucose. Furthermore no evidence was found for the phosphorylytic pathway in mesophyll chloroplasts involving PSH1, while PSH1 was identified in BS chloroplasts very confidently with 57 MS/MS spectra. The maltose exporter MEX1 was 2-fold more abundant in BS chloroplasts, whereas the 23 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. glucose transporter was 2-fold more abundant in mesophyll chloroplasts; both expression patterns are consistent with these different preferences for starch degradation. We identified three ß-amylase homologues, BAM3, BAM6 and BAM9 (Smith et al., 2004; Fulton et al., 2008), respectively type II, I and III. BAM3 and BAM9 were only detected in BS chloroplasts, whereas BAM6 was nearly two-fold higher in M than BS chloroplasts. The functions of maize or Arabidopsis BAM6 and BAM9 have not been studied. Since we did not identify BAM3 in M chloroplasts, it is tempting to speculate that BAM6 represents the major beta-amylose activity in maize mesophyll chloroplasts; it should be noted we only found BAM6 in chloroplast membrane fractions and not in the stroma. The source of starch synthesis in the BS chloroplast begins with condensation of the triose-phosphates DHAP and GAP by fructose-bisphosphate aldolase-2 (SFBA-2) leading to F16BP, followed by dephoshorylation by F16BPase into F6P (in chloroplasts SFBA is both part of the Calvin Cycle and in the dark also glycolysis, whereas F16BPase is unique to the Calvin cycle). In M chloroplasts in maize and non-green plastids in non-photosynthetic tissue, the sources for starch synthesis are triose phosphates (3GPA, possibly directly through the M localized reductive part of the Calvin cycle) and possibly Glc6P imported from the cytosol, mostly likely directly from the BS cells. M chloroplast import of 3PGA from the BS chloroplast occurs through the envelope transporter TPT, whereas it is known that in non-photosynthetic plastids in sink tissues, glucose-6P is imported by the envelope transporter GPT in exchange for TP and/or Pi. We did not observe GPT in the leaf chloroplasts indicating that the source for starch synthesis in M chloroplasts is not imported GlcP, but we observed very significant levels of TPT (in total 245 matched MS/MS spectra). It is therefore most likely that the limited amount of starch produced in M chloroplasts is produced through the reductive phase of the Calvin cycle, followed by the activity of SFBA and F16BP and subsequent conversions by Glc6PI and PGM, similar to BS chloroplasts. We note that PGM2 is mostly localized in the BS chloroplast (BS/M ratio is 3.3), whereas the ten-fold more dominant form PGM1 is equally distributed over both cell types (Fig. 6). The key step in regulation of starch synthesis occurs at the level of AGPase through allosteric regulation by 3PGA (stimulation) and Pi (inhibition) as well as redox regulation, in addition to the availability of ATP and Glc1P (Zeeman et al., 2007). Regulation of starch synthesis is BS chloroplasts most likely will follow the C3-type regulation, which is well-studied 24 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. in Arabidopsis leaves (Zeeman et al., 2007). Regulation of starch synthesis in M chloroplasts is likely rate-limited by the availability to generate sufficient Glc1P. Functional characterization of BAM6 and BAM9, as well as the composition of M chloroplast localized starch are needed to fully understand the role of transient starch storage in C4 leaves. Redox regulation and defense against oxidative stress Redox regulation and defense against radical oxygen species (ROS) is of key importance for optimal functionality of the chloroplast. We identified 45 proteins and protein groups (in total 54 genes) that we classified as being involved in chloroplasts redox regulation and/or oxidative stress defense; most proteins were soluble proteins identified in the stroma (Table 2). Whereas we did identify and quantify some of those in our initial stromal BS-M analysis (Majeran et al., 2005), the newly quantified set is far more exhaustive, but correlates well with our initial observations. Table 2 shows BS/M accumulation ratios, as well as relative protein abundances for each of the annotated proteins. We identified and quantified the known key chloroplast protein components involved in detoxification of superoxide and hydrogen peroxide (H2O2), as well as the glutathione defense system (Table 2). In addition we identified five different glutaredoxins (in total seven genes), four peroxiredoxins and one rubredoxin. The most abundant proteins (based on nSAF) were 2Cys Peroxiredoxin A,B (Prx A,B) (with 4999 MS/MS spectra), Cu,Zn superoxide dismutase, thioredoxin m2&4 and Peroxiredoxin IIE (PrxII E); these were all soluble proteins, identified in the stromal fractions. Except for a low abundant ascorbate peroxidase (33 matched MS/MS spectra), all components of this detoxification system were between 2 and 5-fold enriched in the M chloroplasts; this is consistent with the much higher linear electron transport rates and water splitting activity by PSII and the associated risk of generation of oxygen radicals see (Majeran et al., 2005; Majeran, 2009) for more discussion and references. We identified three FNR proteins (all three with high numbers of MS/MS spectra), as well as 5 ferredoxin (Fd) proteins. These Fd proteins match to Arabidopsis Fd1 (AT1G10960.1), Fd2 (AT1G60950.1), Fd3 (AT2G27510.1), and to an uncharacterized Fd protein (AT4G14890.1). Maize Fd3 was only identified with 5 matched MS/MS spectra, whereas the others were identified with many more spectra, ranging from 44 (Fd1) to 518 (Fd2-1). Fd2-2 was BSenriched, whereas the other were enriched in M chloroplasts. We mention these proteins here 25 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. since the Fd-FNR system connects thylakoid electron transport activity to metabolic activity via NADPH (Table 2). Redox regulation through the thioredoxin system is important in chloroplasts and can activate and deactivate metabolic pathways (Michelet et al., 2006; Lemaire et al., 2007; Schurmann and Buchanan, 2008). We identified and quantified 10 different chloroplast thioredoxins (in total 13 genes). Interestingly, we also identified and quantified the α and ß subunits of the Fd-dependent thioredoxin reductases (FTR-A/B) (Schurmann and Buchanan, 2008), as well as the NADPH-dependent thioredoxin reductase C (NTRC). NTR contains both an NADP-thioredoxin reductase (NTR) and a thioredoxin (Trx) domain and NTRC is able to conjugate both NTR and TRX activities to reduce 2Cys Prx using NADPH as a source of reducing power (rather than Fd). NTRC may also more generally help to integrate regulation of metabolic processes, including regulation of starch synthesis (Lepisto et al., 2009; Michalska et al., 2009). NTRC was very strongly (10-fold) enriched in M chloroplasts. Two SOUL domain proteins, possibly involved in heme delivery or degradation, were also identified. Strikingly, except for thioredoxin (CDSP32), thioredoxin h-2 (Trx-H2), thylakoid bound APX (t-APX)-2 and SOUL heme-binding, that were enriched in the BS chloroplasts, all other proteins were preferentially located in the M chloroplast. NTRC (with 80 matched MS/MS spectra) and two of the glutaredoxins (with 137 and 23 MS/MS spectra) showed the most extreme BS/M ratio of 0.09, 0.08 and 0.05 (Table 2). The differential accumulation of redox and ROS defense components shows that BS and M chloroplast metabolism operates under different redox and oxygen radical stress conditions; Cell-type specific quantitative and qualitative differences in redox regulators and ROS defense components are in place to cope with these different environments. Nucleotide metabolism and homeostasis of nucleotides Nucleotides are critical molecules in nearly every aspect of plant life, and function in primary and secondary metabolism, as well as gene expression. Purine nucleotides (ATP and GTP) and pyrimidine (UTP and CTP) nucleotides are predominantly synthesized in plastids (Zrenner et al., 2006). Their synthesis requires high amounts of energy. Phosphotransfer reactions by kinases and phosphatases convert monoand di-nucleotides to tri-nucleotides and also equilibrate different pools of nucleotides; this is important to balance the activity of different chloroplast 26 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. metabolic pathways and to re-equilibrate between chloroplasts and the cytosol. In addition, conversion of the pyridine nucleotide cofactor NAD or NAD to NADP(H) by NAD(H) kinases is important to accommodate the activity of NADor NADP-specific enzyme activities. Since BS and M chloroplasts in C4 plants have such different roles in photosynthesis and carbon metabolism, we were particularly interested to determine specific adaptations of these two chloroplast-types in terms of nucleotide metabolism and homeostasis. However, this is challenging, since many of the biosynthetic enzymes are know to accumulate at very low levels, and often they are under-represented in proteomics studies. Despite the low abundance of many enzymes, we identified 23 proteins/groups of proteins (in total 33 genes) involved in nucleotide metabolism and homestasis (Fig. 7). All proteins were found in the soluble stromal fraction, with the exception of one of the inorganic pyrophosphatases, adenylate monophosphate kinase 5 (AMK5) and of course the envelope nucleotide translocator (AATP1/NTT1); these were detected in the chloroplast membrane fractions. The most abundant proteins were stromal adenylate monophosphate kinase 2 (AMK2) (with more than 5000 matched MS/MS spectra), nucleoside diphosphate kinase 2 (NDPK2), and soluble inorganic pyrophosphatase. The membrane bound ATP/ADP translocator, AATP1, involved in import of ATP in exchange for ADP, has a BS/M of 0.45 which is consistent with higher photosynthetic electron transport and ATP synthase capacity than in BS cells. We note that we did not detect the plastid envelope localized uniporter BRT1; there are two isoforms (BRT1-1,2) in maize and the Arabidopsis homologue of one of them was recently shown to be involved in export of AMP, ADP and ATP. The other maize isoform serves to export AGPglucose in endosperm. A greater portion of the enzymes are preferentially located in M cells. One possible explanation for this is that de novo biosynthesis of nucleotides is very energy consuming (Zrenner et al., 2006). M chloroplasts are able to generate ATP by linear and cyclic electron flow (Edwards et al., 2001) and may therefore be more capable of driving biosynthesis of nucleotides. However, two enzymes which have high spectral counts, formylglycinamidine ribonucleotide synthase (FGAMS) and CPS, are more concentrated in BS cells (BS/M=3.32 and 3.14 for FGAMS and PYD1 respectively). As discussed in N assimilation, CPS may be responsible for recycling photorespiratory ammonium, thus explaining its preferred localization in BS cells. We have no functional explanation for the preferred BS location of FGAMS. 27 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Fatty acid and lipid metabolism We identified 27 proteins and protein groups (in total 36 genes) involved in fatty acid and lipid metabolism, most of them in the soluble fraction (Fig 8; Suppl. Table 3). 19 enzymes were involved in fatty acid synthesis and elongation, and two stearoyl-ACP desaturase (SSI2/FAB2) and linoleate desaturase (FAD7/FAD8) were involved in fatty acid desaturation. Furthermore, we identified UDP-sulfoquinovose synthase (SQD1) responsible for production of sulfo-lipids. Finally, three enzymes were involved in lipid degradation and 4 proteins were assigned to lipid transport. We were able to annotate and functionally assign most of these proteins and we reconstructed the pathway for fatty acid synthesis (Fig. 8). We did not observe the ACPthioesterases (Fat-A,B) that terminate the elongation reactions; since none of the many proteomics studies in Arabidopsis did identify these either, we conclude that they must have a lower abundance than the other fatty acid synthesis enzymes. Most of the proteins (18 out of 27) were exclusively identified in the soluble stromal fractions; proteins exclusively identified in the membrane fraction were components of the PDH complex and phoshatidic acid-binding protein (TGD2) associated at the inner envelope membrane, two of the lipases, linoleate desaturase (FAD7/FAD8), and a lipocalin domain protein. Protein abundance of the identified proteins (based on nSAF) did span nearly 4 orders of magnitude, with two acyl carrier proteins (ACP3,4) being the most abundant proteins. The high abundance of these carrier proteins is consistent with their function. In our initial stromal analysis (Majeran et al., 2005), we identified only four proteins involved in fatty acid and lipid metabolism; three were preferentially expressed in M chloroplasts. Our current analysis greatly expands the coverage of these pathways, which allowed us to better assess differences between M and BS localized fatty acid and lipid metabolism. The average and median BS/M ratio for all proteins involved in fatty acid synthesis was 0.91 and 0.94, respectively, indicating comparable distribution of fatty acid synthesis across both cell types. This suggests that demand for fatty acids is similar in both BS and M cells. However, we observed clear differential BS-M expression for two of the lipases DAD1 hydrolase (Fig. 8) (see further below) Fatty acids are strictly synthesized in the chloroplast (and non-green plastids) and synthesis occurs by sequential addition of 2-carbon units to acyl groups attached to a soluble acyl 28 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. carrier protein (CAP). Acetyl CoA is the initial carbon precursor, and the building block for elongation. In C3 chloroplasts, most acetyl-CoA is generated from pyruvate by pyruvate dehydrogenase, rather than through acetyl-CoA synthase using imported acetate as source (Fig. 8). Pyruvate was clearly not generated through chloroplast glycolysis, since we did not identify any of the three plastid glycolytic enzymes. Through the C4 carbon cycle, high levels of pyruvate are generated in BS chloroplasts and redistributed to M chloroplasts; this should provide a good resource for synthesis of acetyl-CoA, even if it is in direct competition with the Calvin cycle. The pyruvate pool could be replenished by import from the cytosol, but our data do not allow us to assess that contribution. It was interesting to observe that acetyl-CoA synthetase (also named acetate-CoA ligase) (ACS) was only found in M chloroplasts at significant levels (59 AdjSPC). This provides an alternative route for acetyl-CoA production, independent of pyruvate and PDH. The glycerol precursor phosphatidic acid can be synthesized within the plastid from the Calvin cycle intermediate DHAP through the prokaryotic pathway and involves DHAP-reductase and acyl-ACP-glycerol 3-P-acyltransferase (ATS1,2). Alternatively, phosphatidic acid is imported from the ER via envelop proteins TDG1 and TDG2. Phosphatidic acid is then dephoshorylated by an inner envelope phosphatase (PAP) to produce diacylglycerol (DAG) followed by enzymatic transfer or one or two molecules of galactose resulting in MGDG and DGDG. Alternatively, sulfo-quinovose is transferred from UDP-sulfoquinovose to the DAG portion of PA to produce the sulfo-lipid SQDG. We identified UDP-sulfoquinovose synthase (SQD1), and the subsequent enzyme Fd-glutamate synthetase (GLU1) (see N-assimilation) leading to the production of UDP-sulfoquinovose, but none of the actual enzymes that catalyze the conversion of PA to these various lipid classes (i.e. SQD2, MGD1, DGD1). An indirect alternative source for glycerol is the breakdown of phospatidyl-glycerol (PC) by lipases. We identified three lipases in the chloroplast membrane fractions but their precise function is unknown. Two of these lipases, in particular the very abundant DAD1 hydrolase, showed preferential M accumulation (Fig. 8). The TGD2 gene encodes a phosphatidic acid-binding protein tethered to the inner chloroplast envelope membrane facing the outer envelope membrane. It is proposed that TGD2 represents the substrate-binding or regulatory component of a phosphatidic acid/lipid transport complex in the chloroplast inner envelope membrane. We identified TGD2 with 21 AdjSPC in M membranes but not in BS membranes, suggesting 29 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. differential regulation of lipid transport between chloroplasts and the ER in BS compared to M membranes. An integrative overview of cell-type specific expression patterns of isoprenoid and tetrapyrole metabolism Isoprenoids and tetrapyroles play a central and absolutely essential role in functioning of the chloroplast, and they include tocopherols, quinones, carotenoids, chlorophyll and heme. The enzymes of these pathways are distributed over the soluble and membrane phase, with the upstream steps located in the stroma, and down-stream steps in the chloroplast membranes. Twenty-two 22 proteins involved in the isoprenoid pathway were identified, 20 of which we could assign to specific steps (Fig. 9). We also identified 21 proteins in tetrapyrole synthesis and 2 involved in chlorophyll degradation and this information was integrated with the isoprenoid pathway (Fig. 10). Except for DXS, we indentified all enzymes in the methylerythritol 4phosphate pathway (MEP) and except for Urogen III synthase, Mg-protomethyltransferase and the D subunit of Mg-chelatase, we identified the complete pathway leading to chlorophyllide, including the regulators FLU and GUN4 (Fig. 9). Finally, we also indentified two of the three enzymes (Fe-chelatase and heme oxygenase) of the branch leading to production of phytochromobilin. For some of the enzymes we identified 2 homologues. Finally, VTE1 (tocopherol cyclase in the plastoglobules), VTE3 (MPBQ/MSBQ methyl transferase in the inner envelope) and VTE4 (tocopherol methyltransferase) involved in tocopherol synthesis were also identified. Geranylgeranyl pyrophosphate also serves as the precursor for carotenoid synthesis. We identified six enzymes in carotenoid synthesis, and with the exception of phytoene desaturase, these proteins were of low abundance (Fig. 9). These pathways were integrated in figure 9 and the relative protein concentrations (based on nSAF values) in BS and M chloroplasts are displayed to obtain a unique overview of abundance of the various steps, as well as BS-M chloroplast specific accumulation patterns. Chlorophylls and heme are required in both BS and M chloroplasts, with a higher demand for chlorophyll b in M chloroplasts due to the higher level of PSII light harvesting complexes. The average and median BS/M ratios for chlorophyll and heme pathway proteins were respectively 1.2 and 0.67; this suggests a quite equal distribution across the two cell types, 30 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. possible with some preferential accumulation in M chloroplasts. For discussion of some of the membrane bound enzymes we refer to (Majeran et al., 2008). Isoprenoids can be synthesized in the cytosolic mevalonate (MVA) pathway or the chloroplast-localized MEP pathway (Phillips et al., 2008; Cordoba et al., 2009). IPP is an end product of both pathways and can possibly be transported across the chloroplast envelope. However, it is generally believed that the MEP pathway provides most, if not all, precursors for synthesis of carotenoids, chlorophyll, quinones and tocopherol. The MEP pathway and the downstream enzymes GPPS, GGPS accumulated at higher levels in the M than BS chloroplasts. This is likely due to the much higher demands for carotenoids in the M chloroplasts, as indicated by the low BS/M ratios for several of the enzymes (ZEP, CCD, LUT1, LCY-beta) (Fig. 9). This higher demand for carotenoids must relate to their role in quenching of excess light and detoxification of triplet chlorophyll and singlet oxygen in particular generated during linear electron transport. Similarly, we observed somewhat preferred M accumulation of the VTE enzymes involved in tocopherol and plastoquinone synthesis (Fig. 10); both components are needed in the thylakoid membrane, in particular during linear electron transport. N-assimilation from inorganic and organic N-sources Chloroplasts play a central role in N-assimilation. During nitrogen assimilation, nitrogen is incorporated into amino acids in the form of ammonium (NH4) (Fig. 10; Table 3). Plants obtain ammonium from two sources. The primary source originates from inorganic nitrogen, either by reduction of nitrate (NO3) or as direct uptake of ammonium from soil or symbiotic rhizobium. The secondary source is derived from organic compounds within the plant, through processes such as photorespiration. Upon transport of nitrate through the vascular system into the BS and M cells, nitrate is reduced to nitrite by nitrate reductase (NR) in the cytosol (Lillo, 2008). Nitrite is then imported into the chloroplast by the nitrite transporter (Sugiura et al., 2007), followed by further reduction into ammonia by Fd-nitrite reductase (NiR) and subsequent incorporation of ammonia into glutamine (Gln) in the GOGAT/GS cycle and its redistribution to other amino acids through asparate transaminase. An alternative NH4 assimilation is catalyzed by formation of carbamoylphosphate (CP) for synthesis of arginine (Fig. 11) and purimidine nucleotides (Fig. 8). 31 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. We did not identify the nitrite transporter in maize, mostly likely due to its low abundance; the Arabidopsis homologue has not (yet) been identified in any of the published proteomics studies either. However, we did identify and quantify the major nitrogen assimilation enzymes within chloroplasts (Fig. 10; Table 3): Fd-nitrite reductase (NiR) (BS/M=0.18), glutamine synthase 2 (GS2) (BS/M=1.33), Fd-dependent and NADH-dependent glutamate synthases (Fdand NADH-GOGAT) (BS/M=1.00 and 5.33, respectively). Previous studies have indicated that primary nitrogen assimilation takes place in M cells, with the Fd-NiR and FdGOGAT predominantly localized to M cells while GS activity is present in both cell types (Rathnam and Edwards, 1976; Harel et al., 1977; Becker et al., 1993). Our proteomic data of NiR and GS2 are consistent with these findings. In the case of Fd-GOGAT, we found equal abundance in BS and M cells. The explanation for this apparent discrepancy explanation likely lies in its participation in both primary (preferentially in M) and secondary nitrogen assimilation (preferentially in BS from photorespiration). NADH-GOGAT is strongly BS enriched (BS/M=5.33), but its overall abundance is 100-fold lower (based on nSAF values) than FdGOGAT. NADH-GOGAT is believed to generally express higher in plant roots than in leaves, but unlike Fd-GOGAT, it has not been studied in much detail. The strong preferential accumulation of NADH-GOGAT likely reflects a metabolic adaptation to the BS chloroplast environment. Our results are consistent with the notion that inorganic N assimilation it tightly correlated with photosynthesis and its distribution between BS and M must depend on the availability of reducing equivalents for Fd-NiR activity. We detected three related chloroplast envelope transporters, OMT1 (BS/M 0.09), DiT1 (BS/M = 0.26) and DiT2 (BS/M = 1.16). DiT1 transports 2-OG into the chloroplast, whereas DiT2 exports Glu into the cytosol; both transporters use malate as the counter transport molecule (Fig. 10). The precise role of OMT1 is unclear, but is clearly preferentially accumulating in M chloroplasts, similar to DiT1. OMT1 and DiT1 are strongly (4-10 fold) enriched in the M chloroplast (BS/M=0.09 and 0.26, respectively), likely because the high flux rate of malate from M chloroplasts in the C4 cycle, and low export from BS chloroplasts due to its high rate of malate consumption by ME. Moreover, photorespiration is confined to the BS cells (but operates at lower flux rates than in C3 plants). DiT2 is only slightly higher in BS chloroplasts; using the ZmGI database, we found that DiT2 had a higher BS/M ratio (1.7) (Majeran et al., 2008; 32 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Majeran, 2009). Careful verification of the full length sequences of the DiT family is needed to clarify this difference. Aspartate aminotransferase (AspT) catalyzes the reversible transamination between Glu and oxaloacetate (OAA) to give rise to Asp and 2-oxoglutarate (2-OG). The rate of OAA production in the cytosol (from PEP by PepC) and subsequent import into M chloroplasts is high since this is part of the C4-cycle. AspT is ~2-fold higher in M chloroplasts (BS/M=0.54), consistent with our initial observation (Majeran et al., 2005). The production of Asp does compete with production of malate and must involve several regulatory steps. In addition AspT may provide a link between nitrogen and carbon pathways in maize, if Asp is used to transport carbon to the BS cells, as in NAD-ME and PEPCK types of C4. Carbamoylphosphate synthetase (CPS) condenses ammonium (or the Gln amide group) and HCO3 to make carbamoylphosphate, a precursor for arginine and pyrimidine biosynthesis (Fig. 10). It was indicated that, in C3 plants, mitochondrial CPS is involved in recycling photorespiratory nitrogen (Taira et al., 2004; Potel et al., 2009). Here we observed higher levels of the CPS large subunit-2 (LSU-2) and small subunit (SSU) in BS cells (BS/M=4.16 and 2.43 respectively), while CPS LSU-1 level is comparable (BS/M=1.03). This implies that the chloroplast-targeted CPS may also play a role in recovery of photorespiratory nitrogen. Unlike CPS, most enzymes in the Arg synthesis pathway are preferentially located in M cells (see further in the section on amino acid biosynthesis below); this is best explained by a lower availability of Glu in BS chloroplasts, as Glu is needed in the GOGAT cycle to remove ammonium released from photorespiration. In Arabidopsis, N-assimilation in integrated with carbon metabolism via the chloroplastlocalized nitrogen sensor protein PII (Smith et al., 2003) and it interacts with N-acetyl glutamate kinase (NAGK) (Chen et al., 2006). The PII protein senses concentrations of ATP and 2oxoglutarate and relieves arginine feedback inhibition of NAGK. A similar scenario seems in place in rice (Sugiyama et al., 2004). We identified PII in Arabidopsis chloroplasts with many MS/MS spectra (see PPDB). Surprisingly, there is no obvious maize homologue of the rice or Arabidopsis PII protein, which suggest that integration of carbon and nitrogen metabolism is organized differently in maize and possibly other C4 species. S-assimilation and synthesis of methionine, cysteine and glutathione 33 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Sulfur is absorbed by plants in the form of sulfate, which is first reduced to sulfite (SO3) and then to sulfide (S) before being incorporated into cysteine. Sulfate assimilation requires high amounts of reducing equivalents and therefore occurs mostly in the photosynthetically active leaves; reduced sulfur compounds are then distributed to sink tissues via the phloem (Hopkins et al., 2004; Kopriva and Koprivova, 2005). Sulfate reduction takes place exclusively in chloroplasts, whereas cysteine can be synthesized in chloroplasts, cytosol and mitochondria – see (Krueger et al., 2009) and references therein. But since sulfate reduction takes place exclusively in plastids in both C3 and C4 species, cytosolic or mitochondrial cysteine synthesis is limited by transport of sulfide out of the chloroplast. We obtained good coverage of enzymes involved in the step-wise reduction of sulfate to sulfite and then to sulfide (Fig. 11; Table 3). Coverage of subsequent synthesis of cysteine, methionine and glutathione was more scattered likely relating to the distribution of these synthetic pathways across multiple subcellular compartments (Fig. 12). Older studies using other assays than proteomics revealed that the activities of ATP sulfurylase (ATPS) and adenosine 5’phosphosulfate sulfotransferase (APR) in maize were confined to BS cells, while cysteine synthase (OASTL) activity is found in both cell types but at a higher level in M cells (Gerwick and Black, 1979; Burnell, 1984; Schmutz and Brunold, 1984; Burgener et al., 1998). These previous observations are consistent with our current observations for ATPS2 and APR3,4, which showed BS/M ratios of 3.86, 9.92 and 19.75, respectively, whereas two O-acetylserine thiol lyase isoforms (OASTL) (also named cysteine synthase) were 40-100% higher in M chloroplasts than BS chloroplasts. OASTL was very abundant (identified with 504 MS/MS spectra), but we did not identify serine acetyl Co-A transferase (SAT) which is only active when associated with OASTL. The lack of detectable levels of SAT is consistent with a previous report indicating that OASTL is far more abundant than SAT see (Hopkins et al., 2004)). In addition, SAT activity is also reported to be high in mitochondria and cytosol, and the possibility of OAS transport between subcellular compartments is suggested by experimental data (Krueger et al., 2009). Interestingly, we found that the highly abundant sulfite reductase (SiR; identified with 172 adjSPC) was equally distributed across BS and M chloroplasts (BS/M = 1.08), suggesting that sulfite is transported from BS to M chloroplasts, in addition to well-established cysteine transport (Burgener et al., 1998). Reduction of sulfite into sulfide in M chloroplasts does 34 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. decrease the demand for NADPH in BS, which should be beneficial as BS chloroplasts lack linear photosynthetic electron transport activities. (Fig. 11) Cysteine is used for synthesis of methionine in a three-step reaction, involving CGS, cystathionine β-lyase (CBL) and methionine synthase (MetS); we did not observe CGS, but we found CBL and MetS preferentially located in BS cells (BS/M = 4.72 and 10.01 respectively). CBL clearly appears a bonafide chloroplast localized enzyme (also when comparing to other maize samples – unpublished). However, MetS is quite possibly a contamination from the cytosol. MetS subcellular localization was believed to be cytosolic, but recently it was shown for Arabidopsis that one of the isoforms of MetS (MS3) is localized in the plastid and MetS activity in the plastid was supported by experimental evidence (Hacham et al., 2008). The MetS protein that we identified was very abundant in total maize leaf extract and isolated BS strands, and even if this MetS protein is not with the chloroplast, it is highly enriched in the BS cells, as indicated in figure 11. Cysteine is also the source for S atoms for proteins with Fe-S clusters; these include several enzymes in S-assimilation (APR and SiR), as well as enzymes in N-assimilation (NiR, GOGAT) and photosynthetic electron transport (Ye et al., 2006; Xu and Moller, 2008). We identified eight proteins involved in Fe-S formation: cysteine sulfurase (cpSufS/cpNifS) and its activator cpSufE extract elemental S from cysteine, cpSufB,C,D and three NFU proteins that contribute to Fe-S cluster assembly (Fig. 11; Table 3). These five proteins were on average 3fold enriched in the M chloroplast; given the complexity of sulfur metabolism and its integration with Fehomeostasis, there is no a simple explanation for this M-enrichment. The tripeptide glutathione (GSH) is an important thiol functioning in defense and ROS scavenging (see earlier section on redox and ROS) and GSH is also a source of electrons for the sulfate-reduction (Fig. 11). GSH is synthesized from glutamate and glycine in a two-step reaction, involving γ-glutamylcysteine synthetase (γ-ECS or GSH1) and glutathione synthase. γECS is exclusively localized in plastids in all plant species, whereas GSHS and glutathione synthesis also (or predominantly) occurs in the cytosol (as studied in various C3 species). We detected γ-ECS (with 36 adjSPC) with a larger portion of the total protein located in M chloroplasts (BS/M = 0.66), consistent with feeding experiments that showed preferential GSH synthesis in M cells (Burgener et al., 1998). We did not detect (GSHS) in the BS or M chloroplasts which is consistent with report that it is localized in both cytosol and plastids, thus 35 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. possibly lowering the plastid GSHS concentration (Gomez et al., 2004), but we did detect glutathione reductase (GR) at very high levels (230 adjSPC), responsible for converting oxidized glutathione (GSSG) to reduced glutathione (GSH). GR was three fold higher in M chloroplasts, consistent with the higher rate of ROS production (see section of redox regulation and ROS defense). Our observations clarify several of the controversies for distribution of sulfur metabolism in maize BS and M cells see (Kopriva and Koprivova, 2005). Amino acid metabolism Amino acid synthesis in plants is distributed across chloroplasts, cytosol and mitochondria. Some amino acids (e.g. Gln) act primarily to assimilate and transport nitrogen. Amino acids are also precursors of many nitrogen-containing compounds. Therefore, amino acid biosynthesis is regulated through various mechanisms and with various alternative pathways, and is tightly coordinated with carbon metabolism. Given this complexity, it is perhaps not surprising that very little is known about the distribution of amino acid biosynthetic pathways between BS and M cells in C4 plants. In our previous BS-M stromal analysis we could only identify a handful of enzymes involved in amino acid biosynthesis (Majeran et al., 2005). In our current study, we identified and functionally assigned some 60 proteins/protein groups (about 70 genes) involved in amino acid metabolism (with an overlap to Sand N-assimilation and photorespiration) and they covered 3 orders of magnitude in relative concentration (based on nSAF values) (Table 3). Amino acid biosynthetic pathways can be separated based on the precursor, but connections between these pathways exist. We have grouped the identified proteins in different pathways in table 3, and a number of proteins and pathways are integrated in the figures for N and Sassimilation (eg for Arg and Cys biosynthesis). Aspartate aminotransferase (AspT) was the most abundant protein (based on nSAF; 1687 MS/MS spectra; BS/M =0.54), followed by shikimate kinase SKL1 (488 MS/MS spectra; BS/M =0.7) involved in biosynthesis of aromatic amino acids. Among the proteins with at least 40 adjSC, we observed BS/M ratios ranging from 0.06 (>10-fold enriched in M chloroplast) to 5.96 (6-fold enriched in BS chloroplast), with the majority of proteins being enriched in M chloroplasts (Table 3). In the remainder of this section we will only briefly describe our findings, with an emphasis on the link to C4 metabolism. 36 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. For the glutamate family (Glu, Arg,Pro), we identified 7 out of the 8 enzymes needed for Arg biosynthesis. Arg is synthesized from Glu and CP and is thus closely linked to the GOGAT/GS cycle. All seven identified Arg biosynthetic enzymes were enriched in the M chloroplast (on average 2.5 fold) (Table 3; Fig. 10). There must be competition between Arg and nucleotide synthesis as both require CP as precursor. We suggest that Arg synthesis and accumulation serves to redistribute excess assimilated N to other parts of the plant. It is interesting to note that in the Arabidopsis GLU1 mutant, excess photorespiratory ammonium was transported in the form of Arg (and other amino acids) (Potel et al., 2009). Asp is the precursor for homoSer, Thr, Iso, Lys and also contributes to the synthesis of Met. We identified nine enzymes in these pathways, with particular the Lys synthesis branched being well covered (Table 3). The pathway was M-enriched which is understandable given the high rates of Asp synthesis and high energy requirements (ATP and NADPH). We identified most of the enzymes involved in synthesis of the branched aminoacids, Val and Leu (Table 3). The precursor of this pathway is pyruvate and the last step in their synthesis requires Glu as amine source. The later steps in Leu synthesis are clearly not M-enriched, but either equally distributed across both cell type or enriched in the BS-cell; the reason is not clear. As shown in Fig. 11, we identified several of the enzymes involved in Ser and Cys synthesis (Table 3), as explained in the section on S-assimilation. We did not identify proteins involved in Gly synthesis, which is reassuring since Gly is primarily produced in mitochondria from Thr, Ser or photorespiration. The shikimate pathway for synthesis of the aromatic residues (Phe, Tyr, Trp) was well covered with 11 proteins, even if the number of spectral counts was low for several of the steps (see comment in Conclusions) (Table 3). The entire shikimate pathway takes place in plastids and starts with the condensation of erythrose 4-phosphate (a product of the pentose phosphate pathway and of the Calvin cycle) and PEP, followed by incorporation of a second molecule of PEP in the penultimate step of the central pathway. This central part of the pathway produces chorismate in seven steps, followed by two separate pathways for synthesis of Trp and of Tyr plus Phe. We identified three steps (1, 5 and 6) in the central pathway and multiple steps in the Trp and Tyr/Phe specific pathways. All, but one of the enzymes were more highly expressed in the M chloroplast (the BS-enriched exception was identified by only 4 spectral counts and is therefore not reliable). EPSP synthase (3-phosphoshikimate 1-carboxyvinyltransferase), in the 37 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. central part of the shikimate pathway, is the target of the well-known and extremely ‘popular’ herbicide glyphosate; EPSP synthase was 5-fold enriched in M chloroplasts. The preferential accumulation in M chloroplasts is in line with the high levels of PEP synthesis in the M chloroplast for the C4 carbon cycle. The availability and origin of erythrose 4-phosphate in the M chloroplast is less clear, but must be generated by the pentose phosphate pathway, rather than the Calvin cycle. Indeed, this is consistent with the observed accumulation of OPPP enzymes in the M chloroplast (G6PDH, Lactonate and 6PGDH; see Fig. 5). The last step in Trp synthesis is carried out by subunits of the Trp synthase complex; these subunits were 3-4 fold higher in M than in BS chloroplasts (Table 3). Synthesis of histidine (His) follows a linear pathway, originating with phosphoribosyl pyrophosphate (PRPP) (Fig. 8). We identified enzymes for six out of eight steps of the pathway, even if the number of spectral counts was low for all but the last step, histidinol dehydrogenase (DHD; 59 matched MS/MS spectra). The BS/M ratio of DHD was 0.66 but the distribution of the complete His pathway across both cell types is not clear. Clearly, the His pathway in C4 species required more attention. Many proteins of unknown function show differential BS-M accumulation In addition to the proteins discussed above, we identified and quantified over 200 proteins (and groups) with either a miscellaneous function (e.g. TPR and PPR proteins, rhodanese and DnaJ domain proteins) or without any obvious function (Suppl. Table 3). 96 were only detected in the stromal fractions and 25 were identified in both stroma and membrane fractions. These proteins spanned 3 orders of abundance (based on nSAF) and upto 1556 matched spectra; a significant number of proteins showed strong preferential BS or M accumulation are a an excellent resource for further exploration of specialized BS or M chloroplast functions. CONCLUSIONS Large scale mass spectrometry analysis with a high resolution and high sensitivity mass spectrometer has allowed pathways and activities to be analyzed and reconstructed across the differentiated BS and M chloroplast in the leaf of maize. Quantitative information about relative protein accumulation levels and cell-type specific protein accumulation patterns provided new insight into the functions and structures of differentiated BS and M chloroplasts. These results 38 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. provide a major step forward from previous analyses and also give many new entry points for studying specific aspects of chloroplast biogenesis, differentiation and function. We did note that a number of proteins, particular in the functional categories of nucleotide metabolism and DNA interactions and transcription, were quantified with only few spectra counts. This is likely due to a down-regulation of these functions when chloroplasts begin to reach the end point of biogenesis and differentiation. For instance, non-photosynthetic chloroplast (etioplasts) at the leaf base will be expected to contain much higher protein levels for these functions. The availability of the first maize genome sequence has provided an excellent template for protein identification and quantification, even if imperfections surely must have resulted in missed proteins and inaccuracies of protein quantification. Through this study we have added a wealth of information to these maize protein accession numbers, and this includes assignment of protein names, function and subcellular localization; this will all be made available on-line through the PPDB. From PPDB, this information can be freely distributed to the various maize databases (e.g. http://maizesequence.org/index.html and http://www.maizegdb.org/) and other resources. Moreover, all mass spectral data are made available through the public depository PRIDE (http://www.ebi.ac.uk/pride/). A central conclusion from this study is that differentiated BS chloroplasts at the maize leaf tip appear to have strongly reduced plastid protein expression and protein import. Instead, it is likely actively remodeling it’s proteome, as evidenced by increased levels of ClpB and HSP90. Furthermore, this study provide strong experimental support for a number of specialized BS and M metabolic functions, including starch biosynthesis (BS), Arg synthesis (M), MEP activity (M), N-assimilation (M), initial steps in S-assimilation (BS) and more. Functional and genetic studies, as well as in vitro enzyme activity assays will be needed to further determine their significance. In a forthcoming study, we will explore the protein accumulation of the various pathways and processes along the leaf developmental gradient, and strengthen and expand on our observations presented here for the differentiated BS and M chloroplasts. MATERIALS AND METHODS Maize genotype, plant growth and purification of BS and M chloroplast fractions 39 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. WT-T43 maize plants were grown for 12 to 14 days in a growth chamber (16 h light/8 h dark, 400 μmol photons.m.s) until the 4 leaf was emerging. M and BS chloroplasts were purified from the top 4 cm section of the 3 leaf, harvested about 2 hrs after the onset of the light period, using several hundreds of leaf tips, following procedures as in (Majeran et al., 2005). Purified M and BS chloroplasts were broken with a Dounce homogenizer, and thylakoid and envelope membranes were collected by 20 min centrifugation at 80k xg. The supernatant, representing the enriched soluble stromal cross contamination of M and BS chloroplasts fractions was collected. Cross-contamination was assessed from the presence of the M and BS markers (respectively PPDK and Rubisco) visualized on stained 1-DE SDS-PAGE gels, as described in (Majeran et al., 2005). Protein concentrations were determined with the Bradford essay (Bradford, 1976). Out of more than eight BS-M chloroplast preparations, the best three were selected. Chloroplast stroma and membrane proteome analysis by nanoLC-LTQ-Orbitrap 150 μg of each BS and M stromal preparation were separated by SDS Tricine PAGE (12% acrylamide). Each gel lane was then cut in 12 slices, proteins were digested with trypsin and the extracted peptides were analyzed by nanoLC-LTQ-Orbitrap mass spectrometry using data dependent acquisition and dynamic exclusion, as described in (Majeran et al., 2008). Each sample was analyzed twice with different amounts injected to ensure maximum protein coverage. The complete analysis was carried out in three independent biological replicates. In total 144 MS runs were carried out, with extensive blanks between each sample analysis to avoid carry-over of peptides that could bias quantification. Purification and mass spectrometry analysis of the BS and M membranes is described in (Majeran et al., 2008). Processing of the MS data, database searches and upload into PPDB Peak lists (.mgf format) were generated using DTA supercharge (v1.19) software (http://msquant.sourceforge.net/) and searched with Mascot v2.2 (Matrix Science) against maize genome release 4a.53 (with 53764 models) from http://www.maizesequence.org/ supplemented with the plastid-encoded proteins (111 protein models) and mitochondrial-encoded proteins (165 protein models). For off-line calibration, first a preliminary search was conducted with the precursor tolerance window set at ±30 ppm. Peptides with the ion scores above 40 were chosen as benchmarks to determine the offset for each LC-MS/MS run. This offset was than applied to 40 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. adjust precursor masses in the peak lists of the respective .mgf file for recalibration using a Perl script (unpublished B. Zybailov). The recalibrated peak lists were searched against the maize genome dataset with organellar genes (with 54380 entries) and in parallel against ZmGI v16.0 (with 56364 entries) including sequences for known contaminants (e.g. keratin, trypsin) concatenated with a decoy database where all the sequences were randomized. Each of the peak lists were searched using Mascot v2.2 (maximum p-value of 0.01) for full tryptic peptides using a precursor ion tolerance window set at ±6 ppm, variable methionine oxidation and fixed cysteine carbamido-methylation, and a minimal ion score threshold of 30 for maize genome and 44 for ZmGI; this yielded a peptide false discovery rate (FDR) below 1%, with peptide FPR calculated as: 2*(decoy_hits)/total_hits. The false protein identification rate of protein identified with 2 or more peptides was zero. To reduce false protein identification rate of proteins identified by one peptide, the Mascot search results were further filtered as follows: ion score threshold was increased to 40 for maize genome and 50 for ZmGI, and mass accuracy on the precursor ion was required to be within ±3 ppm. Precursor ion masses below 700 Da were discarded. All filtered results were uploaded into the Plant Proteomics DataBase, PPDB (http://ppdb.tc.cornell.edu/) (Sun et al., 2009). All mass spectral data (the mgf files reformatted as PRIDE XML files) are available via the Proteomics Identifications database (PRIDE) at http://www.ebi.ac.uk/pride/. Selection of the best gene models and post-Mascot filter to assign shared and unique peptides and creation of protein groups with a high percentage of shared matched spectra Many genes have more than 1 gene model, thus leading in many cases to different predicted proteins. In such cases, we selected the protein form (gene model) that had the highest number of matched spectra; if two gene models had the same number of matched spectra, the model with the lowest digit was selected. For quantification by spectral counting, each protein accession was scored for total spectral counts (SPC), unique SPC (uniquely matching to an accession) and adjusted SPC. The latter assigns shared peptides to accessions in proportion to their relative abundance using unique spectral counts for each accession as a basis. Proteins that shared more than ~80% of their matched adjusted peptides with other proteins across the complete dataset were grouped into clusters by generating a similarity matrix through calculation of the dice coefficient between each pair of identified proteins. The dice coefficient is defined as s = 2 |X ∩ 41 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. Y| / ( |x| + |Y| ), where x and y represent the number of SPC for each protein. The similarity cutoff was 0.80. The MCL software (Enright et al., 2002) was used to cluster the proteins into groups, with inflation value set at 5. In some cases, a group contained one protein that had a high percentage of unique SPC (e.g. more than 80%), indicating that this was the most abundant member of the identified protein family. All groups were manually verified and ungrouped if needed. Additional proteins were grouped manually as needed, in particular if they had a low number of adjusted SPC (eg less than 20). The linkage of homologous proteins that were identified by the same set of MS/MS spectra was recorded (their identification marked as ambiguous) and the matched MS/MS spectra were marked as ‘shared’ spectra (or ‘not unique’). For proteins that were indentified with unique spectra, as well as shared spectra, a linkage with the related identified protein was recorded (marked as ‘related’ protein). Calculation of relative abundance and proteins BS/M ratios using nSAF values Relative abundance for each identified protein accession (or cluster) was calculated by the normalized Spectral Abundance Factor (nSAF) (Zybailov et al., 2006) within each technical replicate. SAF was calculated based on the number of adjusted SPC for a protein, normalized by the number of predicted tryptic peptides (for that protein) within a mass range of 700-3500 Da since shorter peptides were excluded from the MASCOT search results, while the longer peptides were beyond the m/z window of MS acquisition. The SAF for each protein was than normalized for the sum of all SAF in the technical replicate, resulting in nSAF. BS/M protein accumulation ratios were calculated based on average nSAF values. Average relative abundance for each protein was calculated for BS membranes, M membranes, BS soluble, M soluble, BS total, M total, and total BS+M. Within the chloroplast, the protein mass in the membraneenriched fractions as compared to the soluble fractions is similar; therefore, to calculate total BS chloroplast and total M relative abundance, average nSAF values for membrane and soluble fractions were summed (Suppl. Tables 2,3). The Plant Proteome Database and functional assignment of identified proteins Mass spectrometry-based information of all identified proteins was extracted from the Mascot search pages and filtered for significance (e.g. minimum ion scores, etc) and ambiguities and shared spectra as described in (Zybailov et al., 2008). This information includes Mowse scores, 42 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. number of matching peptides, number of matched MS/MS spectra (counts), number of unique and adjusted counts, highest peptide score, highest peptide error (in ppm), lowest absolute error (ppm), sequence coverage, tryptic peptide sequences). This information is available in the PPDB (Sun et al., 2009) by using the search function ‘Proteome Experiments’ and selecting the desired output parameters; this search can be restricted to specific experiments. Alternatively, information for specific accessions (either individually or a group) can be extracted using the search function ‘Accessions’ and, if desired, this search can be limited to specific experiments. Finally, information for a particular accession can also be found on each ‘protein report page’. Assignment of protein name of ZmGI and maize genome protein accessions was based on a combination of best BLAST hits in the predicted rice proteome (OsGIv5; from http://rice.plantbiology.msu.edu/), the predicted A. thaliana proteome, ATHv8 (from TAIR http://www.arabidopsis.org/) and our manual annotations for ZmGI v16 (from http://compbio.dfci.harvard.edu/) homologues (Majeran et al., 2005; Majeran et al., 2008; Majeran, 2009). Pair-wise BLAST search results between ATHv8, OSGIv5 and ZMGIv16 are available via PPDB. Each identified proteins was assigned to a molecular function, using the hierarchical, non-redundant classification system developed for MapMan (Thimm et al., 2004) http://gabi.rzpd.de/projects/MapMan/), adjusted after manual verification and information from literature and incorporated into PPDB. Predicted PFAM domains for all predicted maize genome proteins are also available in PPDB. ACKNOWLEDGEMENTS This work was supported by grants to KVW from the National Science Foundation (PGRP0211935 and PGRP0701736). LITERATURE CITED Adam Z, Rudella A, van Wijk KJ (2006) Recent advances in the study of Clp, FtsH and other proteases located in chloroplasts. Curr Opin Plant Biol 9: 234-240. Bantscheff M, Schirle M, Sweetman G, Rick J, Kuster B (2007) Quantitative mass spectrometry in proteomics: a critical review. Anal Bioanal Chem 389: 1017-1031. 43 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. 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PLoS ONE 3: e1994. 52 www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. # proteins* Function M+BS (%) a M (%) b BS (%) c BS/M total d M-membr (%) e BSmembr (%) f BS/M membr g M-sol (%) h BS-sol (%) i BS/M sol j 10 DNA 0.6 0.5 0.7 1.3 1.0 1.4 1.4 0.1 0.0 0.5 21 RNA 1.0 1.1 0.8 0.7 2.2 1.6 0.7 196 Protein expression and fate 10.5 11.4 9.6 0.8 8.3 8.1 1.0 14.4 11.1 0.8 77 protein synthesis (c-encoded) 3.2 4.6 1.9 0.4 3.5 0.2 5.7 3.6 0.6 119 protein sorting, folding, PTM and proteases 7.2 6.8 7.7 1.1 4.8 7.9 1.7 8.7 7.5 0.9 23 Photosystem I 7.9 8.4 7.4 0.9 16.5 14.6 0.9 0.2 0.1 0.5 33 Photosystem II 11.4 15.5 7.3 0.5 26.8 12.7 0.5 4.2 2.0 0.5 5 Cytochrome b6f 1.7 1.6 1.8 1.1 3.3 3.6 1.1 0.0 0.0 0.1 9 ATP-synthase 9.5 7.3 11.7 1.6 13.8 22.9 1.7 0.7 0.4 0.5 30 Cyclic electron flow 6.2 4.1 8.4 2.1 7.7 16.1 2.1 0.4 0.6 1.5 18 Light balance and electron carriers 1.6 2.1 1.1 0.5 2.4 1.4 0.6 1.8 0.8 0.4 25 Calvin cycle & Photorespiration 18.5 13.8 23.3 1.7 2.7 3.8 1.4 24.9 42.9 1.7 5 Rubisco complex & interactants 4.8 2.7 7.0 2.6 0.1 1.0 7.8 5.2 12.9 2.5 6 Calvin cycle reductive phase 8.5 8.7 8.4 1.0 2.4 2.4 1.0 14.9 14.3 1.0 14 rest Calvin cycle & Photorespiration 5.2 2.5 8.0 3.3 0.1 0.4 3.1 4.8 15.6 3.3 7 C4 -malate shuttle + CA and organic acid transfo 8.7 9.4 8.0 0.9 1.4 0.9 0.7 17.5 15.1 0.9 24 starch metabolism 0.9 0.5 1.4 2.8 0.0 0.1 4.2 1.0 2.6 2.7 13 minor CHO metabolism 0.5 0.5 0.6 1.2 0.1 0.2 2.3 0.9 0.9 1.0 8 OPP + 4 glycolysis 0.5 0.4 0.6 1.6 0.8 1.2 1.6 4 N-metabolism 1.2 1.4 1.0 0.8 2.7 2.1 0.8 4 S-assimilation 0.2 0.2 0.2 1.1 0.4 0.5 1.1 51 amino acid metabolism 1.2 1.5 0.8 0.5 3.1 1.7 0.5 12 metal + cofactor-vitamin 0.2 0.2 0.2 0.6 0.5 0.3 0.7 27 lipid metabolism 0.9 0.9 0.9 1.0 0.6 0.5 0.8 1.2 1.3 1.1 24 secondary metabolism 0.8 0.9 0.8 0.9 0.9 1.2 1.4 0.8 0.3 0.4 23 tetrapyrrole synthesis 1.0 1.0 0.9 1.0 0.4 0.7 1.8 1.6 1.2 0.8 22 nucleotide metabolism 1.9 2.1 1.6 0.8 0.1 0.0 0.7 4.2 3.2 0.8 7 hormone metabolism 1.2 1.4 1.0 0.8 0.3 0.7 2.5 2.4 1.4 0.6 37 membrane transport 1.7 1.9 1.6 0.8 3.5 3.0 0.9 0.3 0.1 0.5 37 redox.regulation 3.8 4.8 2.8 0.6 1.5 0.7 0.5 8.1 4.9 0.6 10 stress 0.2 0.1 0.2 1.3 0.2 0.4 1.8 0.1 30 various minor functions 0.7 0.8 0.7 0.9 1.1 1.1 1.0 0.4 0.2 0.5 213 unkown+miscellaneous 5.4 6.2 4.6 0.7 7.5 5.7 0.8 5.0 3.5 0.7 Table 1. Distribution of protein mass (based on % nadjSPC) towards different functions in Mesophyll (M) and Bundle sheath (BS) chloroplast fractions expressed as percentage of the total fraction. * number of protein accessions, after grouping. (a-b-c) Distribution of protein mass (based on % nadjSPC) towards different functions within (a) M + BS together, (b) M alone, (c) BS alone. (d,g,,j) BS/M protein acumulation ratios in (d) membrane + soluble frations, (g) membrane fractions and (j) soluble fractions. (e-f-h-i) Distribution of protein mass (based on % nadjSPC) towards different functions in (e) M membranes alone, (f) BS membranes alone, (h) M soluble fraction alone, and (i) BS soluble fraction alone. www.plantphysiol.org on December 31, 2017 Published by Downloaded from Copyright © 2010 American Society of Plant Biologists. All rights reserved. accession # or group id a lab annotation Curated Loc. Mem/S tr c BS/M d ∑adjSPC e nSAF x 10 f GRMZM2G106928_P01 copper/zinc superoxide dismutase Cu,Zn-SOD (CSD2) S S 0.73 2831 1303.8 GRMZM2G006791_P01 thylakoid bound APX (t-APX) T-i 526 0.41 680 100.4 GRMZM2G120517_P01 ascorbate peroxidase (APX) S S 2.22 36 5.7 GRMZM2G047968_P01 ascorbate peroxidase (APX) S S 0.10 8 2.7 GRMZM2G057709_P02 monodehydroascorbate reductase (MDHAR) S S 0.23 325 42.0 AC202439.3_FGP003 dehydroascorbate reductase-2 (DHAR-2) S S 0.35 271 63.5 GRMZM2G156227_P01 APX-like protein lumen (APX-like) T-lumen 1.37 0.48 526 81.6 GRMZM2G172322_P01 glutathione reductase (GR) S S 0.35 230 28.0 GRMZM2G012479_P01 glutathione peroxidase 2 (GPX2) S 0.09 0.46 338 99.0 72 gamma-glutamylcysteine synthetase (GSH1 or γ-ECS) S S 0.66 36 3.8 100 2-Cys Peroxiredoxin A,B (Prx A,B) S 0.02 0.74 4999 1101.3 GRMZM2G036921_P01 Peroxiredoxin IIE (PrxII E) S 0.02 0.76 1721 478.5 GRMZM2G012276_P01 Peroxiredoxin Q (Prx Q) T-lumen 0.52 0.38 395 154.4 37 glutaredoxin T M 0.10 3 0.6 186 glutaredoxin S S 0.08 137 54.0 GRMZM2G113418_P01 glutaredoxin S S 0.05 23 10.5 GRMZM2G053726_P01 glutaredoxin S S 0.92 150 34.6 GRMZM2G117300_P01 glutaredoxin S S 0.57 88 17.5 GRMZM2G130379_P01 rubredoxin protein T-i M 0.79 142 71.3 GRMZM2G139803_P01 ferredoxin-thioredoxin reductase alpha (FTR-A) S S 0.68 419 145.7 GRMZM2G157458_P01 ferredoxin-thioredoxin reductase alpha (FTR-A) S S 0.80 73 27.6 GRMZM2G026656_P01 ferredoxin-thioredoxin reductase alpha (FTR-A) S S 0.41 23 5.0 GRMZM2G122793_P01 ferredoxin-thioredoxin reductase-beta (FTR-B) S S 0.14 35 12.9 GRMZM2G059083_P01 ferredoxin-NADP(+) reductase 1 (FNR-1) T-p-s 1.43 0.4