reconstruction of metabolic pathways, protein expression ... · 1 this work was supported by the...

Reconstruction of Metabolic Pathways, ProteinExpression, and Homeostasis Machineries acrossMaize Bundle Sheath and Mesophyll Chloroplasts:Large-Scale Quantitative Proteomics Using the FirstMaize Genome Assembly1[W][OA]

Giulia Friso2, Wojciech Majeran2,3, Mingshu Huang, Qi Sun, and Klaas J. van Wijk*

Department of Plant Biology (G.F., W.M., M.H., K.J.v.W.) and Computational Biology Service Unit (Q.S.),Cornell University, Ithaca, New York 14853

Chloroplasts in differentiated bundle sheath (BS) and mesophyll (M) cells of maize (Zea mays) leaves are specialized toaccommodate C4 photosynthesis. This study provides a reconstruction of how metabolic pathways, protein expression, andhomeostasis functions are quantitatively distributed across BS and M chloroplasts. This yielded new insights into cellularspecialization. The experimental analysis was based on high-accuracy mass spectrometry, protein quantification by spectralcounting, and the first maize genome assembly. A bioinformatics workflow was developed to deal with gene models, proteinfamilies, and gene duplications related to the polyploidy of maize; this avoided overidentification of proteins and resulted inmore accurate protein quantification. A total of 1,105 proteins were assigned as potential chloroplast proteins, annotated forfunction, and quantified. Nearly complete coverage of primary carbon, starch, and tetrapyrole metabolism, as well as excellentcoverage for fatty acid synthesis, isoprenoid, sulfur, nitrogen, and amino acid metabolism, was obtained. This showed, forexample, quantitative and qualitative cell type-specific specialization in starch biosynthesis, arginine synthesis, nitrogenassimilation, and initial steps in sulfur assimilation. An extensive overview of BS and M chloroplast protein expression andhomeostasis machineries (more than 200 proteins) demonstrated qualitative and quantitative differences between M and BSchloroplasts and BS-enhanced levels of the specialized chaperones ClpB3 and HSP90 that suggest active remodeling of the BSproteome. The reconstructed pathways are presented as detailed flow diagrams including annotation, relative proteinabundance, and cell-specific expression pattern. Protein annotation and identification data, and projection of matched peptideson the protein models, are available online through the Plant Proteome Database.

Plants can be classified as C3 or C4 species based onthe primary product of carbon fixation in photosyn-thesis. The primary product of carbon fixation is afour-carbon compound (oxaloacetate [OAA]) in C4plants but a three-carbon compound (3-phosphoglyc-erate [3PGA]) in C3 plants. In leaves of C4 grassessuch as maize (Zea mays), photosynthetic activities arepartitioned between two anatomically and biochemi-

cally distinct bundle sheath (BS) and mesophyll (M)cells. A single ring of BS cells surrounds the vascularbundle, followed by a concentric ring of specializedM cells, creating the classical Kranz anatomy. Activecarbon transport (in the form of C4 organic acids)from M cell to BS cells and specific expression ofRubisco in the BS cells allows Rubisco, the carboxyl-ating enzyme in the Calvin cycle, to operate in a highCO2 concentration. The high CO2 concentration sup-presses the oxygenation reaction by Rubisco (and thesubsequent energy-wasteful photorespiratory path-way), resulting in increased photosynthetic yield andmore efficient use of water and nitrogen. The historyof C4 research has been described (Nelson and Langdale,1992; Sage and Monson, 1999; Edwards et al., 2001). Atpresent, there is renewed interest in C4 photosynthe-sis, stimulated in part by the potential use of C4 plantsas a source of biofuels (Carpita and McCann, 2008)and the genetic engineering of C4 rice (Oryza sativa;Sheehy et al., 2007; Hibberd et al., 2008; Taniguchiet al., 2008). The use of new genomics and/or pro-teomics tools has resulted in new insights into cellu-lar differentiation in C4 plants (Majeran and VanWijk,2009).

1 This work was supported by the National Science Foundation(grant nos. PGRP–0211935 and PGRP– 0701736 to K.J.v.W.).

2 These authors contributed equally to the article.3 Present address: Institut des Sciences du Vegetal UPR 2355,

CNRS, Avenue de la Terrasse 23, F–91198 Gif-sur-Yvette cedex,France.

* Corresponding author; e-mail [email protected] author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policydescribed in the Instructions for Authors (www.plantphysiol.org) is:Klaas J. van Wijk ([email protected]).

[W] The online version of this article contains Web-only data.[OA] Open Access articles can be viewed online without a sub-

scription.www.plantphysiol.org/cgi/doi/10.1104/pp.109.152694

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Proteins are responsible for most cellular functions,and knowing their abundance, cell-type specific ex-pression patterns, and subcellular localization is es-sential to understand C4 differentiation. Previously,we published a quantitative analysis of purified Mand BS chloroplast (soluble) stromal proteomes inwhich BS-M protein accumulation ratios for 125 ac-cessions were determined; this covered a limitedrange of plastid functions, although it enabled theintegration of information from previous studies(Majeran et al., 2005). A subsequent complementaryquantitative proteomics study, using nano-liquidchromatography (LC)-LTQ-Orbitrap mass spectrom-etry (MS) and label-free spectral counting comple-mented with other techniques, identified proteins inBS and M thylakoid and envelope membranes ofmaize chloroplasts and determined cell type-specificdifferences in (1) the protein assembly state andcomposition of the four photosynthetic complexesand of a new type of NADPH dehydrogenase (NDH)complex; (2) the auxiliary functions of the thylakoidproteome; and (3) protein and metabolite transportfunctions of M and BS chloroplast envelopes (Majeranet al., 2008). Comparative MS analysis of chloro-plast envelope membranes from leaves of pea (Pisumsativum), a C3 species, and from M chloroplasts ofmaize showed an enrichment of several known andputative translocators in the maize M envelopes(Brautigam et al., 2008). The conclusions of theseproteome analyses are summarized by Majeran andvan Wijk, 2009.

Whereas these proteomics studies provide signifi-cant progress in understanding the organization of C4metabolism in maize, three aspects have not beenadequately addressed: (1) the stromal proteomes of BSandM chloroplasts likely each contain more than 1,500proteins, but the BS-M ratios for only approximately125 proteins were quantified, resulting in very limitedcoverage of several important secondary metabolicpathways such as sulfur, fatty acid, amino acid, andnucleotide metabolism; (2) information about relativeconcentrations of stromal proteins in BS and M chlo-roplasts is lacking but is needed as a basis for quan-titative modeling and metabolic engineering of C4photosynthesis and other metabolic pathways; thegrowing “toolbox” of proteomics and MS now allowsfor such quantitative analyses (Bantscheff et al., 2007;Kumar andMann, 2009); (3) the soluble (Majeran et al.,2005) and membrane (Majeran et al., 2008) proteomedata sets were analyzed by different techniques andmass spectrometers, mostly due to the improvement ofcommercial mass spectrometers in that time frame.Therefore, it is difficult to understand the quantitativerelationships between these data sets. This studyaddresses these three aspects.

So far, maize proteome analyses used essentiallyZmGI maize assemblies (for the Z. mays Gene Index)based on ESTs, combined with a limited amount ofadditional DNA sequence information. The ZmGI wasoriginally generated by The Institute for Genome

Research and subsequently supported by the compu-tational Biology and Functional Genomics Laboratory(http://compbio.dfci.harvard.edu/index.html). ThisZmGI database did not have annotated gene models(for proteome analysis, the DNA sequences weresearched in all six reading frames), and low expressedgenes were likely underrepresented. In our most re-cent BS-M chloroplast analyses (Majeran et al., 2008) aswell as a maize envelope analysis (Brautigam et al.,2008), the MS data were searched against ZmGI ver-sion 16.0 or 17.0. Since that time, the maize genome hasbeen sequenced (using a bacterial artificial chromo-some approach), a physical map was created (themaize accessioned golden path AGP version 1), and itsfirst assembly with gene coordinates and predictedproteins was very recently released (June 2009; http://ftp.maizesequence.org/release-4a.53/sequences/) andpublished (Schnable et al., 2009). This release contains32,540 genes with 53,764 gene models; most of thegene models are evidence based. The new maizegenome assembly is expected to improve maize pro-teome analysis with more accurate protein identifica-tion and quantitative assessment of protein expressionpatterns. This also allows for the determination ofN-terminal localization signals, which was rarely pos-sible from EST assemblies, as N termini were oftenlacking.

This study presents a quantitative protein expres-sion atlas of differentiated maize leaf M and BS chlo-roplasts using high-resolution and mass-accuracy MS(using a LTQ-Orbitrap) and the new maize genomeassembly. Three biological replicates of stromal pro-teomes of isolated BS and M chloroplasts were ana-lyzed. Quantification was carried out based on the“spectral counting” method (Zybailov et al., 2005,2008; Bantscheff et al., 2007; Choi et al., 2008) using asophisticated bioinformatics “workflow” in particularto deal with gene duplications and extended genefamilies observed in polyploids such as maize. Thesenew stromal data sets were combined with a reanal-ysis of our recent BS and M membrane proteome datasets (Majeran et al., 2008) against genome 4a53. Com-pared with previous maize leaf proteome analyses,this study provides an integrated overview of bothprimary and especially secondary metabolism, as wellas chloroplast gene expression and protein biogenesis,in far greater depth. The reconstructed pathways arepresented as figures that include quantitative proteininformation; pathways include primary carbon me-tabolism, starch metabolism, nucleotide metabolism,fatty acid and lipid biosynthesis, chlorophyll, heme,and carotenoid synthesis, and nitrogen assimilation.We briefly comment on the use of the new maizegenome assembly for proteome analysis. All matchedpeptides are projected on the predicted protein modelsvia the Plant Proteomics Database (PPDB; http://ppdb.tc.cornell.edu/). Interactive functional annota-tion, chloroplast localization assignments, as well asdetails of protein identification are also available viaPPDB.

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RESULTS AND DISCUSSION

Purified BS and M Chloroplast Stroma, Analyzed byOne-Dimensional SDS-PAGE and NanoLC-Orbitrap MS

M and BS chloroplasts were purified from the tip ofthe third leaf of 12- to 14-d-old maize plants in whichthe fourth leaf was emerging, as described (Majeranet al., 2005). The soluble stromal proteomes werecollected, selected for purity, and analyzed by one-dimensional SDS-PAGE, followed by in-gel digestionand analysis of extracted peptides by nanoLC-OrbitrapMS. The complete analysis was carried out in threeindependent biological replicates. In total, 144 MSruns were carried out, and all spectral data weresearched against maize genome assembly release 4a53,supplemented with the mitochondrial and chloroplastgenomes. We identified 1,662 protein models corre-sponding to 954 protein accessions when countingonly one model per gene (see below; SupplementalTable S1).

Purified BS and M Chloroplast Membranes, Analyzed byBlue Native-PAGE and NanoLC-Orbitrap MS

Previously, we determined the BS andM chloroplastmembrane proteomes using protein separation bynative gels, followed by tryptic digestion and exten-sive MS/MS analysis by LTQ-Orbitrap (Majeran et al.,2008). Since no genome assembly was available at thattime, we searched the data against the ZmGI assembly(version 16.0). Here, we researched these MS dataagainst the maize genome release 4a53. We identified1,219 protein models corresponding to 882 proteinaccessions when counting only one model per gene(see below; Supplemental Table S1).

Integration of Stromal and Membrane Data Sets andSelection of the Best Gene Model

The workflow of the proteome analysis is summa-rized in Figure 2. Combining the stromal and mem-brane data sets identified 2,439 maize genemodels and1,429 protein accessions when counting only onemodel per gene (Supplemental Table S1). In the newmaize genome annotation, many genes have morethan one gene model. In such cases, we selected theprotein form (gene model) that had the highest num-ber of matched spectra (spectral counts [SPC]) acrossall experiments; if two gene models had the samenumber of matched spectra, the model with the lowestdigit was selected. Within each protein report page inPPDB, “pop-up windows” display the gene modelsfor each protein, with the peptide count projected onthe exons and details for matched peptides projectedon the primary amino acid sequence. This will allowthe user to determine the significance of the differentgene models. BLASTsearch of the 1,429 maize proteinsagainst the Arabidopsis (Arabidopsis thaliana) pro-teome and rice genome resulted in 1,017 Arabidopsis

and 1,079 rice homologues; 47 and 48 are chloroplastencoded in Arabidopsis and rice, respectively.

Relative Amounts and Concentrations of Proteins in BSor M Cells

The relative amount (mass) of each identified pro-tein within each replicate was calculated based onthe adjusted number of matched MS/MS spectra(adjSPC), normalized by the sum of adjusted SPC inthe replicate, yielding nadjSPC. The adjSPC are thesum of unique SPC and a proportional distribution ofshared SPC, using the ratio of unique SPC to determinethis distribution; we previously developed and testedthis strategy for Arabidopsis proteomes (Zybailovet al., 2008). The relative concentration for each iden-tified protein was calculated as the normalized spectralabundance factor (nSAF), which was calculated fromadjSPC weighted for the number of theoretical trypticpeptides with a relevant length (“observable peptides”;Zybailov et al., 2006).

To better understand the distribution and expres-sion range of the identified protein population, wedetermined their frequency distribution of the relativeconcentration using Log10(nSAF) values and a bin sizeof 0.50 (Fig. 1B). Bins in the range of 21.5 to 23, withthe approximately 110 most abundant proteins, werestrongly dominated by proteins involved in photosyn-thesis and the C4 shuttle. Bins in the range of 25.5 to27, with the approximately 205 proteins of lowestabundance, contained mostly proteins with less thanfive matched spectra, frequently with most of thespectra shared with other proteins; thus, this groupof low-abundance proteins is enriched in low ex-pressed members of groups of homologues, possiblyindicating that these are pseudogenes as well as false-positive identifications (see below).

Removal of Low Scoring Proteins

Our workflow has a strictly controlled false-positivepeptide identification rate, and we use high-resolution(100,000), high-mass accuracy (6 ppm) MS data; thisworkflow helps to avoid false-positive identifications.However, the high complexity of the maize genomeand the presence of repetitive DNA and the fact thatwe are using the first draft genome sequence couldlead to identifications of less meaningful protein ac-cessions (e.g. misassemblies, pseudogenes, etc). There-fore, for further analysis of BS and M chloroplastfunctions, we removed those protein accessions withonly one matched MS/MS spectrum (total of 100accessions). In addition, we removed those proteinaccessions that were identified with only one aminoacid sequence (irrespective of charge state and possi-ble posttranslational modifications) if the sequencehad less than 10 amino acid residues and contained Ile(I) or Leu (L); these two residues have the same mass(they are isobaric) and thus cannot be distinguished byMS. These steps removed 149 proteins, mostly repre-

Protein Expression Atlas of BS and M Maize Chloroplasts

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senting pseudogenes, false-positive identifications,and proteins expressed at very low levels; they repre-sented 0.07% of the calculated protein mass (percent-age of total nadSPC; Supplemental Table S2).

Assignment of Chloroplast Localization

Since we were interested in BS and M chloroplastdifferentiation and function, we evaluated the remain-ing identified proteins for chloroplast localization. Forassignment of a protein to the chloroplast, we used acombination of MS-based scores (chloroplast proteins

should generally have higher scores in chloroplast-enriched fractions than in total leaf fractions) andknown localization of the predicted best homologue inArabidopsis or rice, in addition to the known orpredicted maize protein function. We did pay carefulattention to localization assignment for proteins thatare members of groups of closely related identifiedhomologues.

We assigned 974 proteins to the chloroplast, 175proteins were assigned to other subcellular locationsand were considered contaminants, and for the re-maining proteins we could not assign a subcellular

Figure 1. Protein analysis of chloroplast fractions and protein distribution according to function. A, One-dimensional SDS-Tricine-PAGE (12% acrylamide) of stroma proteins from isolated BS and M maize chloroplasts. Three independent BS-Mreplicates are shown; 150 mg of protein was loaded in each lane. B, Frequency distribution of relative protein abundance in BSand M chloroplasts, calculated from Log10(nSAF) values. Each bin on the x axis corresponds to 0.50 orders of magnitude, withthe total protein population spanning about 6 orders of magnitude. Four protein populations are displayed, and they are allidentified proteins (gray bars), proteins assigned a chloroplast location (black bars), proteins with unassigned subcellularlocalization (striped bars), and nonchloroplast proteins (white bars). C, Distribution of proteins with assigned locations tochloroplast or with unclear locations based on the number of proteins (top) or protein mass based on nadjSPC (bottom) overprimary carbon metabolism and photosynthesis (I), secondary metabolism (II), membrane transport (III), miscellaneous andunknown functions (IV), and plastid gene expression and protein homeostasis (V). D, Distribution of proteins with assignedlocations to chloroplast or with unclear locations to the different functions according to the number of proteins. The mainfunctional groups I, II, III, IV, and V are as defined for C.

Friso et al.




location (Supplemental Table S2). These three cate-gories had respectively 99%, 0.6%, and 0.3% of thetotal nadjSPC, indicating that assigned contaminationsmade up less than 0.6% of the total protein mass. Theactual contamination was lower, since highly ex-pressed (chloroplast) proteins are underestimated intheir abundance using the spectral counting technique(Zybailov et al., 2008). We then analyzed the frequencydistribution of relative protein concentration [based onLog10(nSAF)] for these three groups of proteins. Thisshowed that the chloroplast proteins spanned 4 ordersof magnitude, peaking at 23.5 to 24, whereas thecontaminant proteins spanned about 3.5 orders ofmagnitude in abundance, peaking at 25.5 to 26 (Fig.1B). For further analysis, we removed the contaminantproteins.

The Presence of Organellar Genes in the NuclearGenome Assembly

In addition to the nuclear genome, plastid and mi-tochondria also have a genome, and we merged thesesequences with the 4a53 nuclear genes when wesearched the mass spectral data. Indeed, we identified47 chloroplast-encoded proteins and onemitochondria-encoded protein (Supplemental Table S2). However, wenoted that several of these chloroplast-encoded pro-teins had many shared MS/MS spectra matching tomaize 4a53 (nuclear genome) accessions. BLASTsearching of these maize accessions against Arabidop-sis and rice showed they mostly matched to chloro-plast-encoded Arabidopsis and rice proteins. These4a53 genes could represent pseudogenes from frag-ments of plastid genome inserted into the nuclearchromosomes or could result from DNA sequencingof contaminating chloroplast DNA. Examples are theobservation of seven maize 4a53 genome accessions forthe chloroplast-encoded Rubisco large subunit in addi-tion to the chloroplast accession (NP_043033) and fourmaize 4a53 genome accessions for the chloroplast-encoded PSII subunit cytochrome b559a in addition tothe chloroplast accession (NP_043041). For quantifica-tion purposes, we grouped these redundant accessions,as will be described further below.

Functional Annotation of Proteins, Differentiatingbetween C3 and C4 Enzymes and Splicing

The remaining proteins were assigned a functionusing the MapMan bin classification system (Thimmet al., 2004), similar to its use in previous maize studies(Majeran et al., 2005, 2008). Moreover, the MapManclassification system is well integrated in our PPDB forboth Arabidopsis and maize proteome data (Sun et al.,2009). We added a new bin (assigned 1.5 PS.C4 malateshuttle) for proteins involved in the C4 shuttle (e.g.phosphoenolpyruvate [PEP] carboxylase, pyruvatephosphate dikinase [PPDK], malate dehydrogenase[MDH], and PPDK regulatory protein [PPDK-RP]);importantly, based on quantitative BS-M expression

patterns, we were also able to differentiate betweenproteins specialized in the C4 functions as comparedwith non-C4 functions (e.g. PPDK and MDH). Forinstance, we identified two PPDK accessions,GRMZM2G011507 (PPDK-C4) and GRMZM2G097457(PPDK-C3). PPDK-C4 is extremely abundant, with aBS-M ratio of 0.56, whereas less abundant PPDK-C3 isnearly exclusively localized in BS cells (BS-M ratio of9.7). Interestingly, we identified only one maize geneaccession for PPDK-RP (GRMZM2G004880). PPDK-RP is enriched in M chloroplasts (BS-M ratio of 0.28),indicating its specificity toward the regulation of theM-localized C4-type PPDK. There are no obvioushomologues in the 4a53 genome, and indeed onlyone maize PPDK-RP was identified by bioinformaticsanalysis and cloning (Burnell and Chastain, 2006). InArabidopsis, there are two genes for PPDK-RP (RP1and RP2, At4g21210 and At3g01200, respectively). RP1is similar to the maize C4-type PPDK-RP and is plastidlocalized, whereas RP2 has no detectable activity andis localized to the cytosol (Chastain et al., 2008). Formore details on C4 and non-C4 NADP-MDH enzymes,see Maurino et al. (2001) and Tausta et al. (2002).

The Identified Proteome Provides High Coverage ofSecondary Metabolism, Plastid Gene Expression, andChloroplast Protein Homeostasis

Figure 1C shows the distribution of the number ofproteins (total of 1,105 proteins) or protein mass in-volved in (1) primary carbon metabolism and photo-synthesis, (2) secondary metabolism, (3) miscellaneousand unknown functions, (4) plastid gene expressionand protein homeostasis, and (5) membrane transport.Each of the first four groups involved 23% to 25% ofthe identified proteins, whereas about 4% of identifiedproteins were involved in transport of metabolitesacross the chloroplast envelopes (Fig. 1C, top). Interms of protein biomass, the majority (approximately66%) was invested in primary metabolism, with equaldistribution across secondary metabolism and plastidprotein homeostasis (each 13%; Fig. 1C, bottom).

Amore detailed breakdown of chloroplast functionsis shown in Figure 1D, which shows high coverage offunctions not much observed in maize chloroplasts inprevious studies. In particular, the protein synthesiscomponents involved in expression of chloroplast-encoded proteins were very well covered; this in-cluded most of the ribosomal proteins, many of thet-RNA synthetases, as well as initiation and elongationfactors (Fig. 1D). Even more rewarding was the iden-tification of 132 proteins involved in various aspects ofposttranslational chloroplast protein homeostasis, in-cluding about 50 processing peptidases, aminopepti-dases, and proteases, as well as soluble methionesulfoxide reductases involved in protein repair, pep-tide deformylase, as well as several phosphatases andkinases. Most if not all members of the Clp proteasesystem in maize were identified; while they arenow 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 regula-tors and radical oxygen species (ROS) detoxificationenzymes (e.g. peroxiredoxins), nitrogen, sulfur, andamino acid metabolism, and other secondary meta-bolic pathways, including fatty acid synthesis andisoprenoid, tetrapyrole, and nucleotide metabolism(Fig. 1D). Whereas only a very small percentage ofprotein mass was invested in isoprenoid synthesis andderived products, as well as cofactor and vitaminbiosynthesis (less than 1%), we obtained a good cov-erage of several of these pathways; for instance, mostenzymes of the methylerythritol phosphate pathway(MEP) and multiple enzymes in thiamine (vitamin b1)and riboflavin (vitamin b2) synthesis were identified.Thus, the combined analysis of membrane and solublefractions of isolated BS and M chloroplasts provided agood coverage of many of the chloroplast functions.Therefore, the quantitative comparison of the BS andM profiles should thus provide meaningful and newinsights into C4-driven differentiation and fulfill ourobjective to obtain an integrated, quantitative over-view of the differentiated state of BS andM chloroplastfunctions in the maize leaf.

The Differentiated Functional State of BS and

M Chloroplasts

Table I compares protein mass investment (based onnadjSPC) of BS and M chloroplasts in the differentchloroplast functions (Table I). About 39% (M) to 38%(BS) of the total chloroplast proteome was invested inthe photosynthetic apparatus and cyclic electron flowlocated in the thylakoid membrane (Table I). Between23% (M) and 31% (BS) was invested in the Calvincycle, the C4 shuttle, and carbonic anhydrases. About11% (M) and 10% (BS) was invested in plastid geneexpression, protein folding, processing, and proteoly-sis. About 5% (M) and 3% (BS) was invested in redoxregulation. About 1.9% (M) to 1.6% (BS) was investedin transport functions. Some 6% (M) and 5% (BS) wasinvested in proteins for which the function is un-known. About 14% (M) and 11% (BS) was invested inall remaining functions (Table I). To better appreciatethe cell type-specific differences in molecular func-tions, we further broke down these protein invest-ments for the membrane and soluble fractions (Table I;for a detailed description that also highlights the mostabundant proteins for the various functions, see Sup-plemental Text S1).

Grouping of Identified Protein Accessions to Deal withGene Duplications and Extended Gene Families in Maizefor Quantification

Maize is a polyploid with a far larger genome size(2,800 Mb) than Arabidopsis (130 Mb) and rice (430Mb), and maize has a significantly larger amount ofrepetitive sequences (Schnable et al., 2009). The com-plexity of the maize genome and its predicted pro-

teome requires an extra effort for maize proteomics, inparticular to deal with closely related identified pro-teins. These closely related proteins could representtrue duplicated genes, pseudogenes, or could be arti-facts of the genome assembly.

To deal with this complexity of the maize genomefor quantification of BS and M protein expression, wegrouped proteins that shared more than approxi-mately 80% of their matched SPC. Grouping wasdone by generation of a similarity matrix throughcalculation of the dice coefficient between each pair ofidentified proteins based on matched SPC, followedby clustering of the proteins using MCL software(Enright et al., 2002), followed by manual evaluation(see “Materials and Methods”). In total, 313 proteinswere placed in 131 groups (Fig. 2). This groupingavoids overinterpretation of differences observed be-tween M and BS chloroplast proteomes; this strategywas also very useful for Arabidopsis (Rutschow et al.,2008; Zybailov et al., 2009). The “relationships” be-tween proteins that shared matched MS/MS spectraare visible through PPDB. Quantification of proteinswith a low number of adjSPC within a replicate (i.e.below approximately 10 adjSPC) is generally lessaccurate (Kim et al., 2009; Zybailov et al., 2009). Atotal of 378 proteins had less than 40 adjSPC, 179 hadbetween 40 and 100 adjSPC, and 366 had more than100 adjSPC (Fig. 2), and we generally consider thesethree groups as low-, medium-, and high-confidencequantifications, respectively (Supplemental Table S3).

In the remaining sections, metabolic pathways arereconstructed, and the distribution of specific proteinsin the various processes and metabolic pathways overthe BS and M chloroplasts is discussed. We will firstdiscuss the chloroplast gene expression and proteinhomeostasis machineries to understand the assemblyand maintenance of the differentiated chloroplastproteomes. Furthermore, we focus on those metabolicpathways that were not (or were poorly) covered inprevious maize studies, in particular starch metabo-lism, nucleotide metabolism, sulfur and nitrogenassimilation, and isoprenoid and tetrapyrole metabo-lism. We will also analyze the distribution of relativeconcentrations of the proteins, as this will be valuablefor future modeling of metabolic fluxes. In these var-ious plots and analyses, we will use nSAF values, asthis is the best approximation of relative protein con-centration, as well as BS-M accumulation ratios. Thedata will be presented as figures for plastid geneexpression, protein synthesis, and homeostasis, thyla-koid light reactions (Supplemental Fig. S1), envelopetransporters, primary carbon metabolism, starch me-tabolism, nucleotide metabolism, fatty acid and lipidbiosynthesis, chlorophyll, heme, and carotenoid syn-thesis, nitrogen assimilation, and sulfur assimilation,as well as tables for redox regulation and ROS defenseand for amino acidmetabolism.More details regardingaccession numbers, groupings of accessions, MapManbin numbers, nSAF values, matched MS/MS spectra,and more are available in Supplemental Table S3.

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Plastid Gene Expression, Protein Synthesis,

and Homeostasis

The contribution of plastid gene expression andprotein homeostasis on control of BS and M differen-tiation is entirely unclear. For instance, levels of chlo-roplast-encoded (and nucleus-encoded) PSII subunitsare lower in BS than in M chloroplasts, whereaschloroplast-encoded NDH subunits are higher in BSchloroplasts. However, it is not clear if levels of thesechloroplast-encoded proteins are regulated throughcontrol of transcription (or even chromosome copy

number), mRNA processing and stability, translation,or posttranslationally through proteolysis. Further-more, levels of imported nucleus-encoded proteinswithin the BS or M chloroplast could be controlledthrough proteolysis. Both chloroplast- and nucleus-encoded proteins share intraplastid 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 homeo-stasis, assembled in five categories, namely proteinsynthesis (77 proteins), proteolysis and processing (42

Table I. Distribution of protein mass (based on percentage of nadjSPC) toward different functions in M and BS chloroplast fractions expressedas a percentage of the total fraction





proteins), import and posttranslational modifications(34 proteins), folding (38 proteins), and RNA-DNAinteraction (31 proteins). Figure 3 displays the BS-Mratio (based on nSAF), the number of matched adjSPC,and relative abundance of proteins (as a color scale) inthese various functional classes. A total of 114 proteinswere only found in the stroma, whereas 72 were foundin both membrane and stromal fractions, but with awide range of distribution (i.e. between 3,000-foldenriched in the membrane fraction and 100-fold en-riched in the stromal fraction). The systematic quan-titative comparison of BS and M chloroplast proteinsin these processes did identify several strongly differ-entially expressed proteins (markedwith an asterisk inFig. 3); these may contribute to BS and M specializa-tion. We will highlight selected candidate proteins,with emphasis on soluble proteins, and comment onfunctional implications and follow-up analyses (fordetails, see Supplemental Table S3).

Interactors with the Plastid Chromosome

There is very little understanding of how plastidchromosome copy number and organization influencetranscription and contribute to BS-M chloroplast dif-ferentiation. Therefore, we were pleased to identify

eight DNA-interacting proteins (pTAC or nucleoidproteins) and two DNA-repair enzymes. The abundancelevels of these proteins spanned more than 2 ordersof magnitude (based on nSAF), with two membrane-bound nucleoid-associated proteins, pTAC16 andMFP1, being by far the most abundant, with 1,679and 656 matched MS/MS spectra, respectively.pTAC16 of unknown function was 2-fold enriched inthe M, whereas MFP1 was 2-fold enriched in the BSchloroplasts. MFP1 is a coiled-coil DNA-binding pro-tein and is believed to play a role in anchoring theplastid DNA to the envelope and thylakoid mem-brane; its expression is tightly correlated with theaccumulation of thylakoid membranes (Jeong et al.,2003). Two soluble putative DNA-repair enzymes,auvrB/uvrC motif-containing protein and a deoxyri-bodipyrimidine photolyase, follow in abundance andhave BS-M ratios of 0.51 and 0.99, respectively. Thenext three abundant proteins were homologues ofTCP34, pTAC5, and pTAC17; TCP34 and pTAC5 ho-mologues were only found in the membrane fractions,and pTAC17 was mostly found in the soluble phase.The Arabidopsis homologues of these pTAC proteinswere all identified in a highly enriched plastid chro-mosome preparation, but their precise function is notclear (Pfalz et al., 2006; Weber et al., 2006). Targeted

Figure 2. Workflow of the proteome analysis. Details are explained in the text. The inset graph shows the number of matchedadjSPC per identified protein or protein group. A total of 378 proteins were identifiedwith less than 40 adjSPC, 179 proteins wereidentified with between 40 and 100 adjSPC, and 366 proteins were identified with more than 100 adjSPC.

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Figure 3. Distribution of the chloroplast plastid protein expression and protein biogenesis and homeostasis machinery across BSand M chloroplasts. Data include annotations for function and subchloroplast location, BS-M ratios (based on nSAF), relativeprotein accumulation levels (RA) based on nSAF values for the total BS and M chloroplast proteomes and using a color scale, andtotal number of adjSPC.





nucleoid and functional analyses will be needed todetermine the contribution of transcriptional regula-tion to chloroplast BS and M differentiation.

Regulation of Transcription and RNA Metabolism

We identified 21 proteins in this category, includingsix RRM domain proteins, and four ribonucleases inthe soluble stromal fractions covering 2 to 3 orders ofexpression, with the number of matched MS/MSspectra ranging from 1,400 to just 3 (Fig. 3). The mostabundant proteins were homologues of CP33, CP31,CP29, CSP41A, and CSP41B. Homologues for severalof these proteins have been characterized in Arabi-dopsis; examples are RRM protein CP31A (Tillichet al., 2009) and CSP41 (Beligni and Mayfield, 2008;Bollenbach et al., 2009). CSP41B-2 (BS-M = 2.5) andCSP41A (BS-M = 2.1) and the 3# to 5# exoribonucleaseRIF10 (BS-M = 4.9) and a DUF740 protein (BS-M = 3.5)were all higher expressed in BS chloroplasts, whereasseveral other proteins, such as two SET domain pro-teins and a S1 RNA-binding protein, were much moreexpressed in M chloroplasts (BS-M = 0.2, 0.1, and 0.06).

The Chloroplast Translational Machinery

We quantified 76 proteins (and protein groups) thatare part of the chloroplast translational machinery,including 18 tRNA synthetases, two initiation factors(IF2 and IF3) and five elongation factors (BipA, TU, G,and P types), a ribosome-recycling factor, and a pep-tide chain-release factor, as well as 14 subunits of the30S ribosomal particle, 32 subunits of the 50S particle,and three plastid-specific ribosomal proteins (PSRP1,-2, and -3). Relative protein concentrations spanned3 orders of magnitude, with the most abundant pro-tein being EF-TU-1, with both elongation and chap-erone functions (3,163 matched MS/MS spectra).Chloroplast ribosome levels were about 3-fold higherin M chloroplasts than BS chloroplasts (median BS-Mratio for individual subunits was 0.32). Interestingly,the initiation and elongation factors weremore equallydistributed across BS and M, with the exception of thetypA/BipA elongation factor-like protein. TypA/BipAEF is a specialized ribosome-associated translationfactor (Wang et al., 2008) suggested to be required instress responses; interestingly, we observed BipA EF tobe strongly induced in Arabidopsis chloroplast Clpprotease mutants (Kim et al., 2009; Zybailov et al.,2009). The 18 tRNA synthetases were higher in Mchloroplasts, with a median BS-M ratio of 0.14. Over-all, these data suggest higher translational activity inM chloroplasts than BS chloroplasts, particularly at thethylakoid surface.

When comparing the stromal and membrane pro-teomes, we calculated that nearly 40% of ribosomeprotein mass (based on nadjSPC) was found in themembrane fractions. Interestingly, the M membranefractions contain 20-fold more ribosomal protein thanthe BS membranes, which is very striking when com-

pared with the approximately 2-fold difference be-tween the M and BS stroma. Chloroplast ribosomesare known to associate with the thylakoid membranes;indeed, many chloroplast-encoded thylakoid proteinsare synthesized at the membrane surface and cotransla-tionally inserted (Margulies and Michaels, 1975; Kleinet al., 1988; van Wijk et al., 1996; Rohl and van Wijk,2001). We suggest that the very low BS-M ratio ofthylakoid-bound ribosomes reflects a much higher de-mand for synthesis of thylakoid proteins in M chloro-plasts, most likely due to high M abundance of PSIIsubunits and the relative short lifetime of PSII reactioncenter proteins due to light-induced damage. The BS-enriched NDH complex also contains many chloro-plast-encoded proteins, but the relative concentration ofthe NDH complex, and likely also the turnover rate, isseveral fold lower than the PSII complex, thus contrib-uting much less to overall chloroplast translation.

Protein Processing and Proteolysis

We quantified 42 proteases and processing pepti-dases, 26 of which were quantified with medium orhigh confidence. These include 22 stromal proteases,six thylakoid lumen proteins, nine integral thylakoidproteases, and three proteases that we assigned to theinner envelope membrane. As most membrane pro-teins were discovered and discussed previously whensearching the ZmGI database (Majeran et al., 2008), wefocus here on the 22 soluble proteases and peptidases.Twelve of the soluble proteases were members of theClp protease system, and based on a multialignmentanalysis with the Arabidopsis Clp family, we anno-tated the maize Clp proteins. The Clp protease systemis the most abundant soluble protease in Arabidopsischloroplasts (Peltier et al., 2004) and in pea etioplasts(Kanervo et al., 2008) and consists of a proteolytictetradecameric barrel-structured ClpPR complexto which two small ClpT subunits tightly associate(Peltier et al., 2004). ClpC1 and -C2 chaperones areassumed to deliver substrate to the core proteasecomplex (Adam et al., 2006). We were surprised tofind that the most abundant ClpPR subunit in maizechloroplasts was a homologue of Arabidopsis mito-chondrial ClpP2. Clearly, this maize ClpP2 protein isnot mitochondrial, and this finding warrants a morein-depth phylogenetic analysis of the Clp proteasefamily in plants. The average and median BS-M ratioof the Clp system was 0.8 and 0.7, respectively; thisrather equal distribution across BS and M chloroplastsis consistent with the idea that the Clp system servesas a general housekeeping protease, unlike some of theproteases with narrow functions and extreme BS-Mratios, such as SPPA. The slight bias (25%–40%) towardM accumulation may relate to specific control functionsby the Clp protease system of plastid gene expression;this is certainly worth investigating further.

Other abundant proteases were Prep1 (likely in-volved in degradation of cleaved cTPs), several ami-

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nopeptidases (eucyl, glycyl, glutamyl, M24), and a Lonprotease (LON2). Several of these soluble proteasesshowed a strong preferential distribution for either celltype; in particular, glutamyl endopeptidase (cGEP)and abundant glycyl aminopeptidase (M1) were 4-foldand 2-fold higher, respectively, in the BS stroma,whereas stromal DegP2, the very abundant eucylaminopeptidase LAP1, and the M24 aminopeptidaseAPP2 were 3-, 4-, and 5-fold higher, respectively, in Mchloroplasts. The substrates of these proteases areunknown, but the preferential accumulation in onecell type suggests a narrow set of substrates.

Protein Sorting

We identified the major soluble and membranecomponents that control protein import, sorting, andtranslocation in the thylakoid (Tic110/40/55, SRP43/54, SecA/Y, TatC, and HCF106). Tic110, Tic40, SecY,and SecAwere each identified with more than 50 SPC,and their BS-M ratio could be determined with confi-dence. The envelope import components, as well asboth subunits of the SRP particle and soluble SecA,were clearly higher in M chloroplasts, whereas SecYwas equally distributed. Since most of the SecA-dependent abundant lumenal proteins are compo-nents of M-enriched PSII complex, the strong Maccumulation is logical. SecY is the general thylakoidimport channel for membrane proteins, and its equaldistribution across the two cell types is consistent withits general role.

Protein Assembly Factors

We also identified 13 soluble and membrane pro-teins involved in the assembly of thylakoid complexes;these included several factors that function for veryspecific complexes (e.g. for PSII, HCF136 and LPA1;for PSI, PYG7 and YCF4) and at least three factorsinvolved in biogenesis of 2Fe-2S complexes (NFU1,2,3;see also the section on sulfur assimilation below).Factors for PSII-specific complexes were several-foldhigher in M chloroplasts, and those involved in PSIbiogenesis were 30% to 40% higher in BS (Majeranet al., 2008). Thus, the BS-M accumulation ratios of thePSI- and PSII-specific assembly factors correlate wellwith the BS-M ratios of PSI and PSII themselves,indicating that expression of these assembly factorsmust be well coordinated with demand. Strikingly, themaize homologue of Arabidopsis stromal proteinHCF101 (quantified with 69 adjSPC) involved in bio-genesis of 4Fe-4S clusters (not 2Fe-2S) in PSI (Stockeland Oelmuller, 2004), as well as ferredoxin (Fd)-thio-redoxin reductase (FTR; Lezhneva et al., 2004), showed10-fold higher M accumulation. Unlike PSI, FTR ismore highly expressed in M chloroplast than in BSchloroplasts, and further studies on the regulation ofcell-specific expression and accumulation of HCF101may elucidate regulatory networks for chloroplastbiogenesis and differentiation.

Protein (Un)folding and Maturation

We quantified 38 proteins involved in (un)foldingand maturation; none had predicted transmembranedomains. These proteins included lumenal proteinisomerases, general high-abundance stromal chap-erones (several HSP70s and their GrpE nucleotide-exchange factors and CPN60/20/10 proteins), as wellas low-abundance factors (e.g. BSD2 implicated inRubisco assembly). As discussed previously (Majeranet al., 2008), several thylakoid lumen isomerasesshowed distinct preferential accumulation in BS or Mthylakoids, suggesting specific adaptation or sub-strates. General chaperones were quite equally distrib-uted, consistent with their broad array of substrates.Clear exceptions were stromal chaperone HSP90 andClpB3, involved in protein maturation and unfolding,respectively; they were 2- to 6-fold higher in BS chlo-roplasts. Considering the much lower chloroplasttranslation rates in BS chloroplasts, this is surprisingand potentially very important for understanding theBS-M differentiation pathways; we speculate that thisrelates specifically to the specialization of the BS chlo-roplast (see “Conclusion”).

Posttranslational Modifiers

Nucleus-encoded chloroplast proteins can be mod-ified within the chloroplast after import, whereaschloroplast-encoded proteins can be modified duringand after synthesis. We identified two different typesof protein-repair proteins, Met sulfoxide type A4 andribulosamine/erythrulosamine 3-kinase; both wereonly slightly more abundant in M chloroplasts, pos-sibly because the antioxidative systems within thechloroplast have sufficient capacity to prevent pro-tein damage. We identified peptide deformylase 1A(PDF1a), involved in cotranslational removal of theN-terminal formyl group of Met, with a BS-M ratio of0.68; this higher M accumulation is again consistentwith higher translation rates in M chloroplasts. Thefunction of the very abundant and soluble methyl-transferase (246 MS/MS spectra) is unknown, and ithas a similar BS-M ratio as PDF1a. Phosphorylationdoes play an important role in chloroplast metabolismand adaptation, but more systematic studies on stro-mal kinases and phosphatases are now just beginningto emerge (in Arabidopsis; Schliebner et al., 2008;Reiland et al., 2009). We identified two protein phos-phatases 2C (PP2C), a protein Tyr phosphatase and aputative protein kinase inhibitor. The most abundantPP2C (246 MS/MS spectra) was more than 5-foldenriched in BS chloroplasts, whereas the zinc-bindingdomain protein and putative kinase inhibitor (68 MS/MS spectra) were more than 5-fold enriched in Mchloroplasts. Reversible phosphorylation, in additionto redox regulation (see below), plays an importantrole in regulation of plant metabolism, and the studyof BS and M chloroplast-specific (de)phosphorylationnetworks will be an important complement to this study.





The Photochemical Apparatus in the

Thylakoid Membrane

We obtained extensive coverage (121 proteins andprotein groups) of the linear electron transport chain,including PSII (33 proteins), PSI (23 proteins), thecytochrome b6 f complex (five proteins), the ATP syn-thase (nine proteins), lumenal plastocyanin, five Fdproteins, three FNR1 isoforms, and six light stressproteins with chlorophyll-binding domains of the Ohpand Lil families (Supplemental Fig. S1). We also iden-tified cyclic electron flow components of the NDHcomplex and NDH-specific biogenesis factors (23 pro-teins), NDH-independent cyclic electron flow compo-nents of the PGR complex (three PGRL1 and two PGR5homologues), as well as alternative thylakoid terminaloxidases (PTOX or IMMUTANS; two homologues)and PIFI. We note that homologues of the new NDHsubunits that we discovered previously in maize(Majeran et al., 2008) have now also been identifiedin Arabidopsis chloroplasts (Peng et al., 2009). Finally,we identified two state transition kinases (Stn7and Stn8), the thylakoid phosphoprotein TSP9, andthe Ca2+ phosphoprotein (Supplemental Fig. S1,DE-P). Most of these proteins were also identifiedbased on our previous search against the ZmGI ESTassembly database (Majeran et al., 2008); therefore, wewill not discuss these proteins any further. BS-M ratiosfor these proteins per complex or function are shownin Supplemental Figure S1.

Envelope Transporters with Known andUnknown Functions

From our recent BS-M membrane study (Majeranet al., 2008) and general literature, combined withfunctional suggestions from the study of Weber andcolleagues (Brautigam et al., 2008), we (tentatively)assigned functions for 26 chloroplast envelope trans-porters (Majeran and van Wijk, 2009); these includedM-enriched transporters MEP1,2,3,4 with unknownsubstrates, members of the DIT family (Dit1, Dit2, andOMT1), phosphate/triosephosphate translocators(TPT and PPT), the maltose exporter MEX1, ATP/ADP translocator AATP1 or NTT1, and anion trans-porter ANTR2. A number of envelope transporters(e.g. various porins, ATP-binding cassette trans-porters) could not be assigned to specific functions.Researching our membrane data against the maizegenome identified 46 transporters (52 proteins in 46groups). We summarize this information for thesetransporters with (putative) substrates in Figure 4.Log2 BS-M ratios are displayed in small bar diagrams,with the bars shown in black, dark gray, and light gray,which indicate proteins identified with high (morethan 100), medium (40–100), and low (less than 40)numbers of adjSPC, respectively. Proteins that arequantified with at least 40 adjSPC and enriched morethan 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 proteinabundance (based on nSAF) across both cell types isshown as colored squares.

The transporter with the highest relative abundancewas MEP1-1 (equally distributed between BS and M),followed by MEP3/4 (M enriched), PPT (strong Menriched), and TPT (M enriched). Some of these trans-porters are also integrated with information of specificpathways, in particular with primary carbon metabo-lism and the C4-malate shuttle (Fig. 5), starch metab-olism (Fig. 6), and nucleotide and nitrogen metabolism(Figs. 7 and 9). These assignments are based on pre-vious literature, and in the case of the Calvin cycleintermediates (3PGA and dihydroxyacetone phos-phate [DHAP]) and malate shuttle substrates (PEPand OAA), these assignments are speculative. Identi-fication of substrates for many of these transportersshould be of highest priority.

The Calvin Cycle, the C4 Malate Shuttle, the OxidativePentose Phosphate Pathway, and Glycolysis

Whereas our previous stromal analysis, employingtwo-dimensional gels and quadrupole time of flight-based MS analysis, identified several of the enzymesinvolved in the Calvin cycle and the C4 malate shuttle(Majeran et al., 2005), the new stromal analysis usingspectral counting, the LTQ-Orbitrap, and the newmaize genome gives a nearly complete overview.Figure 5 shows an integrated overview with BS-Mratio of the Calvin cycle, the C4 shuttle (PPDK, PPDK-RP, MDH, ME), the (irreversible) oxidative pentosephosphate pathway (G6PDH, Lact, 6PGDH) and thereversible pentose phosphate pathway (TKL, RPE,RPI, TA), and enzymes leading to starch biosynthesis(PGM1, PGM2, Glc6PI). Furthermore, Figure 5 showsC3 forms of abundant C4 shuttle enzymes (NADP-MHD-C3 and PPDK-C3). The inset shows the relativeabundance of proteins for both cell types combined.We did not find evidence for accumulation of thespecific chloroplast glycolytic enzymes (PyrK, ENI,PGlyM), nor did we find evidence for PPI-PFK or ATP-PFK involved in the conversion of F6P to F16BP. Wedid identify PGP, one of the two chloroplast-localizedenzymes involved in photorespiration, only in the BSchloroplast. Chloroplast glycerate kinase, involved inthe conversion of glycerate imported from peroxi-somes into 3PGA, is likely GRMZM2G054663, but wenever identified it.

In agreement with our initial analysis in 2005(Majeran et al., 2005), our data strongly suggest thatthe reductive phase of the Calvin cycle, represented byGAPDHB and TPI, is enriched in the M chloroplast.The third enzyme in the reductive phase, PGK1, wasalso slightly higher in the M chloroplast, in contrast tothe homologue PGK2, which was much higher in theBS chloroplast. Moreover, two of the reversible pen-tose phosphate pathway enzymes (RPI and TA;marked with asterisks) are also (somewhat) higher inM chloroplasts. The three enzymes of the oxidative

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PPP (G6PDH, Lact, and 6PGDH) are more expressedin the M chloroplasts. A second low-abundance iso-form of Lact was higher in BS chloroplasts, but thesignificance is unclear, since it was only quantified byeight MS/MS spectra. The increased levels of theOPPP enzymes in M chloroplasts suggest that carbo-hydrates imported from the BS cell feed the OPPP inMchloroplasts, likely as a source of carbon intermediatesfor various pathways. Alternatively, the OPPP path-way may also be higher to provide precursors to theshikimate pathway; indeed, as discussed further be-low (Table III), most enzymes in the shikimate path-way were more abundant in M chloroplasts than inBS chloroplast. The preferential BS accumulation ofPGM1, PGM2, and Glc6PI is not a reflection of in-

creased rates of glycolysis but instead reflects higherrates of starch synthesis in BS chloroplasts (see nextsection). The C3-type PPDK is strongly enriched in BSchloroplasts, whereas C3-type NADP-MDH is equallydistributed across both cell types.

Pathways for Starch Metabolism Show StrongQuantitative and Qualitative Differences between BSand M Chloroplasts

Expression and distribution of starch metabolic en-zymes in C4 leaves have not been systematically stud-ied. We identified and quantified the relativeabundances of 21 chloroplast-localized enzymes in-volved in starch synthesis and degradation. These 21

Figure 4. BS-M protein accumulation ratios and relative protein accumulation levels of transport proteins in the inner and outerenvelopes of BS andM chloroplasts. The relative accumulation levels (based on nSAF) averaged across BS andM chloroplasts areshown as colored squares, with the scale indicated. The BS-M ratios are indicated as small bar diagrams using a log2 scale. Barsfor proteins with at least 80 adjSPC are shown in black, proteins with between 40 and 80 adjSPC are shown in dark gray, andproteins with less then 40 adjSPC are shown in light gray. Protein names are shown in red if the BS-M ratio is greater than 1.5 andif they had at least 40 adjSPC. Protein names are shown in blue if the BS-M ratio is less than 0.67 and if they had at least 40adjSPC. Substrates for proteins are shown. Abbreviations are explained in the text and in Supplemental Table S3.





proteins were assigned names, functions, and posi-tions in the starch metabolic pathway, based on BLASTalignments and extensive literature analysis (Smithet al., 2005; Zeeman et al., 2007; Fulton et al., 2008; Fig.6). We used the nomenclature developed for Arabi-dopsis (Smith et al., 2004). For functional interpreta-tion, we connected the starch pathway to enzymes(PGM1,2 and Glc6PI) involved in conversion betweenGlc1P and Glc6P and fructoses (F16BP and F6P), aswell as the maltose and Glc transporters (MEX1 andGlcT1; Fig. 7). BS-M protein ratios are indicated as bardiagrams, and the total protein abundance (based onnSAF) across both cell types is shown as coloredsquares. The majority of enzymes are more abundantin BS chloroplasts than in M chloroplasts, and the totalinvestment of enzymes in starch metabolism is nearly3-fold higher in BS than in M chloroplasts (Table I);

this is consistent with several previous observations(Spilatro and Preiss, 1987; Lunn and Furbank, 1997;Majeran et al., 2005, 2008) and with the much higherpresence of starch particles in BS chloroplasts than inM chloroplasts.

ADP-Glc-pyrophosphorylase (AGPase), a hetero-tetramer of large and small subunits, is the firstcommitted step in starch biosynthesis and is con-trolled by a combination of allosteric control by 3PGAand inorganic phosphate, redox regulation, and tre-halose (Kolbe et al., 2005; Zeeman et al., 2007). Weidentified two isoforms of the large subunit ofAGPase (AGPL1 and -2) and one isoform for the smallsubunit (AGPS). APGL1 is nearly 10-fold more abun-dant than APGL2; APGL1 is about 2-fold higher in BSthan in M chloroplasts, whereas APGL2 is likely aspecific isoform adapted to M chloroplast conditions,

Figure 5. BS andM chloroplast distribution of proteins involved in primary carbonmetabolism. Proteins include the Calvin cycleand photorespiration, the OPP pathway, proteins leading up to starch synthesis, the malate shuttle, as well as C3 homologues ofmalate shuttle proteins (PPDK andMDH). Tentative transporters are indicated, and assignment is based onMajeran and vanWijk(2009). BS-M bar diagrams and color coding are as described in the legend to Figure 4. Relative accumulation levels (based onnSAF) of proteins averaged across BS and M chloroplasts are shown as colored squares, with the scale indicated. Abbreviationsare explained in the text and in Supplemental Table S3.

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since it is more than 10-fold higher in M than in BSchloroplasts. APGS was approximately 2-fold higher inBS than in M chloroplasts and likely serves both largeisoforms.The product of AGPase, ADP-Glc, is used by a

family of starch synthases (SS) to generate linear Glcpolymers named amylose and branched Glc polymersnamed amylopectin. We identified four starch SS,namely granule-bound SS (GSS), which is needed forsynthesis of long linear chains of amylose, and SSI,SSIIa, and SSIIIb, which generate the linear chains ofamylopectin. GSS was 4.4-fold higher in BS than in Mchloroplasts, consistent with a higher production ofstarch in BS and the requirement of GSS for amylosesynthesis. It is thought that SSI is needed upstream ofSSII and that SSII operates upstream of SSIII; SSI isprimarily responsible for the synthesis of short chains,and SSII and SSIII lengthen these chains further. SSIwas equally distributed between BS and M chloro-plasts, whereas the more abundant SSIIa was morethan 2-fold higher in BS chloroplasts. The low-abun-dance SSIIIb was 1.4-fold higher in BS chloroplasts.

Together, this strongly suggests that starch synthe-sized in M chloroplasts has shorter amylase chains ascompared with B chloroplasts.

We identified three starch (de)branching enzymes(BEIIb1 and -2 and ISA2) involved in the synthesis ofbranched glycans. BEIIb introduces a-1,6 branchpoints and is a so-called class II branching enzyme.Absence of class II BE results in the production of onlylong-chain glucans and no amylopectin. ISA2 is a typeof debranching enzyme involved in synthesis (ratherthan degradation), and it has been suggested that ISA2has a more regulatory function (for discussion, seeZeeman et al., 2007). BEIIb2 was about 100 times moreabundant than BEIIb1 and ISA2. BEIIb2 was more than6-fold enriched in BS chloroplasts, whereas BEIIb1 andISA2 were not detected in M chloroplasts. Together,these findings suggest that the starch produced in BSchloroplasts is more highly branched than starch in Mchloroplasts.

We identified two kinases (PWD and GWD) thatcatalyze the phosphorylation of a glucosyl residue ofamylopectin; these two enzymes stimulate starch deg-

Figure 6. Construction of the starch synthesis and degradation pathways across BS and M chloroplasts. BS-M bar diagrams andcolor coding are as described in the legend to Figure 4. Relative accumulation levels (based on nSAF) of proteins averaged acrossBS and M chloroplasts are shown as colored squares, with the scale indicated. Abbreviations are explained in the text and inSupplemental Table S3.





radation, possibly by loosening the surface of thegranule structure and/or increasing the solubility ofstarch (Smith et al., 2005; Zeeman et al., 2007). Bothhave a similar relative concentration, and both wereenriched (3.5- to 5-fold) in BS chloroplasts; these BS-enriched levels are indicative of and/or consistentwith the reduced accumulation of amylopectin in Mchloroplasts (see below). We also identified the maizehomologue of Arabidopsis phosphoglucan phospha-tase DSP4 (or SEX4; Kotting et al., 2009); DSP4 was3-fold enriched in M chloroplasts. DSP4 phosphory-lation activity is needed to allow effective cleavagehydrolysis of glucans by BAM3 and ISA3. The prefer-ential M accumulation of DSP4 is puzzling and sug-gests a specific adaptation of starch degradation.

There are several starch breakdown pathways inleaves, and they result in maltose, Glc, or Glc1P(Zeeman et al., 2007). We were able to assign maizehomologues to each of these pathways in BS chloro-plasts. In C3 leaves such as Arabidopsis, the maltose isthe main breakdown product and export product oftransient starch in leaves during the dark period.

Based on the relatively high levels of ISA3 as com-pared with DPE1 (approximately 15-fold higher), it islikely that BS chloroplasts also use maltose as the mainexport product in the night. However, our observa-tions of high levels of DPE1 (comparable to ISA3 in BSchloroplasts) and its low BS-M ratio (Fig. 6) suggestthat in M chloroplasts the dominant end product forstarch degradation is Glc. Furthermore, no evidencewas found for the phosphorylytic pathway in Mchloroplasts involving PSH1, while PSH1 was identi-fied in BS chloroplasts very confidently with 57 MS/MS spectra. The maltose exporter MEX1 was 2-foldmore abundant in BS chloroplasts, whereas the Glctransporter was 2-fold more abundant in M chloro-plasts; both expression patterns are consistent withthese different preferences for starch degradation.We identified three b-amylase homologues, BAM3,BAM6, and BAM9 (Smith et al., 2004; Fulton et al.,2008), types II, I, and III, respectively. BAM3 andBAM9 were only detected in BS chloroplasts, whereasBAM6 was nearly 2-fold higher in M than in BSchloroplasts. The functions of maize or Arabidopsis

Figure 7. BS andM chloroplast distribution of proteins involved in purine and pyrimidine nucleotidemetabolism and nucleotidehomeostasis. BS-M bar diagrams and color coding are as described in the legend to Figure 4. Relative accumulation levels (basedon nSAF) of proteins averaged across BS and M chloroplasts are shown as colored squares, with the scale indicated.Abbreviations are explained in the text and in Supplemental Table S3.

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BAM6 and BAM9 have not been studied. Since we didnot identify BAM3 in M chloroplasts, it is tempting tospeculate that BAM6 represents the major b-amyloseactivity in maize M chloroplasts; it should be notedthat we only found BAM6 in chloroplast membranefractions and not in the stroma (possibly in associationwith starch granules).The source of starch synthesis in the BS chloroplast

begins with condensation of the triose phosphatesDHAP and GAP by Fru-bisP aldolase-2 (SFBA-2),leading to F16BP, followed by dephosphorylation byF16BPase into F6P (in chloroplasts, SFBA is part ofboth the Calvin Cycle and in the dark also glycolysis,whereas F16BPase is unique to the Calvin cycle). In Mchloroplasts, the possible sources for starch synthesisare the triose phosphate 3GPA directly imported fromthe BS cells, DHAP generated within the M chloro-plasts through the reductive part of the Calvin cycle,and Glc6P imported from the cytosol and supplied bythe BS cells. M chloroplast import of 3PGA from the BSchloroplast occurs through the envelope transporterTPT, whereas it is known that in nonphotosyntheticplastids in sink tissues, Glc6P is imported by theenvelope transporter GPT in exchange for TP and/orinorganic phosphate. We did not observe GPT in theleaf chloroplasts, indicating that the source for starchsynthesis in M chloroplasts is not imported GlcP, butwe observed very significant levels of TPT (245matched MS/MS spectra). Therefore, it is most likelythat the limited amount of starch produced in Mchloroplasts is produced through the reductive phaseof the Calvin cycle, followed by the activity of SFBAand F16BP and subsequent conversions by Glc6PI andPGM, similar to BS chloroplasts. We note that PGM2 ismostly localized in the BS chloroplast (BS-M ratio is3.3), whereas the 10-fold more dominant form PGM1is equally distributed over both cell types (Fig. 6).The key step in the regulation of starch synthesis

occurs at the level of AGPase through allosteric regu-lation by 3PGA (stimulation) and inorganic phosphate(inhibition) as well as redox regulation, in addition tothe availability of ATP and Glc1P (Zeeman et al., 2007).Regulation of starch synthesis in BS chloroplasts mostlikely will follow the C3-type regulation, which is wellstudied in Arabidopsis leaves (Zeeman et al., 2007).Regulation of starch synthesis inM chloroplasts is likelyrate limited by the availability to generate sufficientGlc1P. Functional characterization of BAM6 and BAM9,as well as the composition of M chloroplast-localizedstarch, are needed to fully understand the role oftransient starch storage in C4 leaves.

Redox Regulation and Defense against Oxidative Stress

Redox regulation and defense against ROS are ofkey importance for optimal functionality of the chlo-roplast. We identified 45 proteins and protein groups(total of 54 genes) that we classified as being involvedin chloroplast redox regulation and/or oxidative stressdefense; most proteins were soluble proteins identified

in the stroma (Table II). Whereas we did identify andquantify some of those in our initial stromal BS-Manalysis (Majeran et al., 2005), the newly quantified setis far more exhaustive, but it correlates well with ourinitial observations. Table II shows BS-M accumulationratios as well as relative protein abundance for each ofthe annotated proteins.

We identified and quantified the known key chlo-roplast protein components involved in detoxificationof superoxide and hydrogen peroxide as well as theglutathione defense system (Table II). In addition, weidentified five different glutaredoxins (total of sevengenes), four peroxiredoxins, and one rubredoxin. Themost abundant proteins (based on nSAF) were 2-CysperoxiredoxinA,B (with 4,999MS/MS spectra), copper/zinc-superoxide dismutase, thioredoxins m2 and m4,and peroxiredoxin IIE; these were all soluble proteins,identified in the stromal fractions. Except for a low-abundance ascorbate peroxidase (33 matched MS/MS spectra), all components of this detoxificationsystem were between 2- and 5-fold enriched in theM chloroplasts; this is consistent with the much higherlinear electron transport rates and water-splitting ac-tivity by PSII and the associated risk of generation ofoxygen radicals (for more discussion and references,see Majeran et al., 2005; Majeran and van Wijk, 2009).

We identified three FNR proteins (all three with highnumbers of MS/MS spectra) as well as five Fdproteins. These Fd proteins match to ArabidopsisFd1 (AT1G10960.1), Fd2 (AT1G60950.1), and Fd3(AT2G27510.1) and to an uncharacterized Fd protein(AT4G14890.1). Maize Fd3 was only identified withfive matchedMS/MS spectra, whereas the others wereidentified with many more spectra, ranging from 44(Fd1) to 518 (Fd2-1). Fd2-2 was BS enriched, whereasthe others were enriched in M chloroplasts. We men-tion these proteins here since the Fd-FNR systemconnects thylakoid electron transport activity to met-abolic activity via NADPH (Table II).

Redox regulation through the thioredoxin systemis important in chloroplasts and can activate anddeactivate metabolic pathways (Michelet et al., 2006;Lemaire et al., 2007; Schurmann and Buchanan, 2008).We identified and quantified 10 different chloroplastthioredoxins (total of 13 genes). Interestingly, we alsoidentified and quantified the a- and b-subunits ofthe Fd-dependent thioredoxin reductases (FTR-A/B;Schurmann and Buchanan, 2008) as well as theNADPH-dependent thioredoxin reductase C (NTRC).NTR contains both an NADP-thioredoxin reductaseand a thioredoxin domain, and NTRC is able to conju-gate both NTR and thioredoxin activities to reduce2Cys peroxiredoxin using NADPH as a source ofreducing power (rather than Fd). NTRC may alsomore generally help to integrate the regulation ofmetabolic processes, including regulation of starchsynthesis (Lepisto et al., 2009; Michalska et al., 2009).NTRC was very strongly (10-fold) enriched in M chlo-roplasts. Two SOUL domain proteins, possibly in-volved in heme delivery or degradation, were also





identified. Strikingly, except for thioredoxin (CDSP32),thioredoxin h-2 (Trx-H2), thylakoid-bound APX-2,and SOUL heme binding, which were enriched in theBS chloroplasts, all other proteins were preferentiallylocated in the M chloroplast. NTRC (with 80 matchedMS/MS spectra) and two of the glutaredoxins (with137 and 23 MS/MS spectra) showed the most extremeBS-M ratios of 0.09, 0.08, and 0.05 (Table II).

The differential accumulation of redox and ROSdefense components shows that BS and M chloroplast

metabolism operates under different redox and oxy-gen radical stress conditions. Cell type-specific quan-titative and qualitative differences in redox regulatorsand ROS defense components are in place to cope withthese different environments.

Nucleotide Metabolism and Homeostasis of Nucleotides

Nucleotides are critical molecules in nearly everyaspect of plant life and function in primary and

Table II. Distribution of redox regulators and the ROS defense machinery across BS and M chloroplasts

Data include annotations for function and subchloroplast location, membrane-stroma ratios (based on nadjSPC) and BS-M ratios (based onnadjSPC), number of matched adjSPC, and relative protein accumulation levels (based on nSAF).

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secondary metabolism as well as gene expression.Purine nucleotides (ATP and GTP) and pyrimidinenucleotides (UTP and CTP) are predominantly syn-thesized in plastids (Zrenner et al., 2006). Theirsynthesis requires high amounts of energy. Phos-photransfer reactions by kinases and phosphatasesconvert mononucleotides and dinucleotides to trinu-cleotides and also equilibrate different pools of nucle-otides; this is important to balance the activity ofdifferent chloroplast metabolic pathways and to re-equilibrate between chloroplasts and the cytosol. Inaddition, conversion of the pyridine nucleotide cofac-tor NAD or NAD+ to NADP(H) by NAD(H) kinases isimportant to accommodate the activity of NAD- orNADP-specific enzyme activities. Since BS and Mchloroplasts in C4 plants have such different roles inphotosynthesis and carbon metabolism, we were par-ticularly interested to determine specific adaptationsof these two chloroplast types in terms of nucleotidemetabolism and homeostasis. However, this is chal-lenging, since many of the biosynthetic enzymes areknown to accumulate at very low levels, and oftenthey are underrepresented in proteomics studies.Despite the low abundance of many enzymes, we

identified 23 proteins/groups of proteins (total of 33genes) involved in nucleotide metabolism and homeo-stasis (Fig. 7). All proteins were found in the solublestromal fraction, with the exception of one of theinorganic pyrophosphatases, adenylate monophos-phate kinase 5, and the envelope nucleotide trans-locator (AATP1/NTT1); these were detected in thechloroplast membrane fractions. The most abundantproteins were stromal adenylate monophosphate ki-nase 2 (with more than 5,000 matched MS/MS spec-tra), nucleoside diphosphate kinase 2, and solubleinorganic pyrophosphatase. The membrane-boundATP/ADP translocator, AATP1, involved in importof ATP in exchange for ADP, has a BS-M ratio of 0.45,which is consistent with higher photosynthetic elec-tron transport and ATP synthase capacity than in BSchloroplasts. We note that we did not detect the plastidenvelope-localized uniporter BRT1; there are two iso-forms (BRT1-1 and -2) in maize, and the Arabidopsishomologue of one of them was recently shown to beinvolved in the export of AMP, ADP, and ATP. Theother maize isoform serves to export AGP-Glc inendosperm.A greater portion of the enzymes in nucleotide

metabolism are preferentially located in M cells. Onepossible explanation for this is that de novo biosynthe-sis of nucleotides is very energy consuming (Zrenneret al., 2006). M chloroplasts are able to generate ATP bylinear and cyclic electron flow (Edwards et al., 2001)and therefore may be more capable of driving biosyn-thesis of nucleotides. However, two enzymes that havehigh SPC, formylglycinamidine ribonucleotide syn-thase (FGAMS) and carbamoylphosphate synthetase(CPS), are more concentrated in BS cells (BS-M = 3.32and 3.14 for FGAMS and CPS, respectively). As dis-cussed in nitrogen assimilation, CPS may be responsi-

ble for recycling photorespiratory ammonium, thusexplaining its preferred localization in BS cells. Wehave no functional explanation for the preferred BSlocation of FGAMS.

Fatty Acid and Lipid Metabolism

We identified 27 proteins and protein groups (totalof 36 genes) involved in fatty acid and lipid metabo-lism, most of them in the soluble fraction (Fig. 8;Supplemental Table S3). Nineteen enzymes were in-volved in fatty acid synthesis and elongation, and twostearoyl-ACP desaturases (SSI2/FAB2) and linoleatedesaturases (FAD7/FAD8) were involved in fattyacid desaturation. Furthermore, we identified UDP-sulfoquinovose synthase (SQD1), which is responsiblefor the production of sulfolipids. Finally, three en-zymes were involved in lipid degradation and fourproteins were assigned to lipid transport. We wereable to annotate and functionally assign most of theseproteins, and we reconstructed the pathway for fattyacid synthesis (Fig. 8). We did not observe the ACP-thioesterases (Fat-A and -B) that terminate the elonga-tion reactions; since none of the many proteomicsstudies in Arabidopsis identified these either, weconclude that they must have a lower abundancethan the other fatty acid synthesis enzymes. Most ofthe proteins (18 out of 27) were exclusively identifiedin the soluble stromal fractions; proteins exclusivelyidentified in the membrane fraction were componentsof the pyruvate dehydrogenase (PDH) complex andphosphatidic acid-binding protein (TGD2) associatedat the inner envelope membrane, two of the lipases,linoleate desaturases (FAD7/FAD8), and a lipocalindomain protein. Protein abundance of the identifiedproteins (based on nSAF) spanned nearly 4 ordersof magnitude, with two acyl carrier proteins (ACP3and -4) being the most abundant proteins. The highabundance of these carrier proteins is consistent withtheir functions.

In our initial stromal analysis (Majeran et al., 2005),we identified only four proteins involved in fatty acidand lipid metabolism; three were preferentially ex-pressed in M chloroplasts. Our analysis here greatlyexpands the coverage of these pathways, which al-lowed us to better assess differences between M- andBS-localized fatty acid and lipid metabolism. The av-erage and median BS-M ratios for all proteins involvedin fatty acid synthesis were 0.91 and 0.94, respectively,indicating comparable distribution of fatty acid syn-thesis across both cell types. This suggests that de-mand for fatty acids is similar in both BS and M cells.However, we observed clear differential BS-M expres-sion for two of the lipases (Fig. 8; see below).

Fatty acids are strictly synthesized in the chloroplast(and nongreen plastids), and synthesis occurs by se-quential addition of two carbon units to acyl groupsattached to a soluble acyl carrier protein (CAP). Acetyl-CoA is the initial carbon precursor and the buildingblock for elongation. In C3 chloroplasts, most acetyl-





CoA is generated from pyruvate by pyruvate dehydro-genase, rather than through acetyl-CoA synthase usingimported acetate as a source (Fig. 8). Pyruvate wasclearly not generated through chloroplast glycolysis,since we did not identify any of the three plastidglycolytic enzymes. Through the C4 carbon cycle, highlevels of pyruvate are generated in BS chloroplasts andredistributed to M chloroplasts; this should provide agood resource for the synthesis of acetyl-CoA, even ifit is in direct competition with the Calvin cycle. Thepyruvate pool could be replenished by import fromthe cytosol, but our data do not allow us to assess thatcontribution. It was interesting that acetyl-CoA syn-thetase (also named acetate-CoA ligase) was onlyfound in M chloroplasts at significant levels (59adjSPC). This provides an alternative route for acetyl-CoA production, independent of pyruvate and pyru-vate dehydrogenase.

The glycerol precursor phosphatidic acid can besynthesized within the plastid from the Calvin cycleintermediate DHAP through the prokaryotic pathwayand involves DHAP reductase and acyl-ACP-glycerol3-phosphate acyltransferase (ATS1 and -2). Alterna-tively, phosphatidic acid is imported from the endo-

plasmic reticulum via envelop proteins TDG1 andTDG2. Phosphatidic acid is then dephosphorylatedby an inner envelope phosphatase (PAP) to pro-duce diacylglycerol followed by enzymatic transferor one or two molecules of Gal, resulting in monoga-lactosyldiacylglycerol and digalactosyldiacylglycerol.Alternatively, sulfoquinovose is transferred from UDP-sulfoquinovose to the diacylglycerol portion of phos-phatidic acid to produce the sulfo-lipid SQDG. Weidentified SQD1 and the subsequent enzyme Fd-Glusynthetase (GLU1; see nitrogen assimilation), leadingto the production of UDP-sulfoquinovose but none ofthe actual enzymes that catalyze the conversion ofphosphatidic acid to these various lipid classes (i.e.SQD2, MGD1, DGD1). An indirect alternative sourcefor glycerol is the breakdown of phosphatidylglycerolby lipases. We identified three lipases in the chloro-plast membrane fractions, but their precise functionsare unknown. Two of these lipases, in particular thevery abundant DAD1 hydrolase, showed preferentialM accumulation (Fig. 8). The TGD2 gene encodes aphosphatidic acid-binding protein tethered to theinner chloroplast envelope membrane facing theouter envelope membrane. It is proposed that TGD2

Figure 8. BS and M chloroplast distribution of proteins involved in fatty acid and lipid metabolism. BS-M bar diagrams and colorcoding are as described in the legend to Figure 4. Abbreviations are explained in the text and in Supplemental Table S3.

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represents the substrate-binding or regulatory com-ponent of a phosphatidic acid/lipid transport com-plex in the chloroplast inner envelope membrane. Weidentified TGD2 with 21 adjSPC in M membranes butnot in BS membranes, suggesting differential regula-tion of lipid transport between chloroplasts and theendoplasmic reticulum in BS compared with Mmem-branes.

An Integrative Overview of Cell Type-Specific

Expression Patterns of Isoprenoid andTetrapyrole Metabolism

Isoprenoids and tetrapyroles play a central andabsolutely essential role in functioning of the chloro-plast, and they include tocopherols, quinones, carot-enoids, chlorophyll, and heme. The enzymes of thesepathways are distributed over the soluble and mem-brane phases, with the upstream steps located in thestroma and the downstream steps in the chloroplastmembranes. Twenty-two proteins involved in theisoprenoid pathway were identified, 20 of which wecould assign to specific steps (Fig. 9). We also identi-fied 21 proteins in tetrapyrole synthesis and twoinvolved in chlorophyll degradation, and this infor-mation was integrated with the isoprenoid pathway(Fig. 10). Except for 1-deoxy-D-xylulose-5-phosphatesynthase, we identified all enzymes in the MEP path-way, and except for urogen III synthase, magnesiumprotomethyltransferase, and the D-subunit of magne-sium chelatase, we identified the complete pathwayleading to chlorophyllide, including the regulatorsFLU and GUN4 (Fig. 9). Finally, we also identifiedtwo of the three enzymes (iron chelatase and hemeoxygenase) of the branch leading to the productionof phytochromobilin. For some of the enzymes, weidentified two homologues. Finally, VTE1 (tocopherolcyclase in the plastoglobules), VTE3 (MPBQ/MSBQmethyltransferase in the inner envelope), and VTE4(tocopherol methyltransferase involved in tocopherolsynthesis) were also identified. Geranylgeranyl pyro-phosphate also serves as the precursor for carotenoidsynthesis. We identified six enzymes in carotenoidsynthesis, and with the exception of phytoene desat-urase, 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 aunique overview of abundance of the various steps, aswell as BS-M chloroplast-specific accumulation pat-terns.Chlorophylls and heme are required in both BS and

M chloroplasts, with a higher demand for chlorophyllb in M chloroplasts due to the higher level of PSII light-harvesting complexes. The average and median BS-Mratios for chlorophyll and heme pathway proteinswere 1.2 and 0.67, respectively; this suggests a quiteequal distribution across the two cell types, possiblywith some preferential accumulation in M chloro-

plasts. For discussion of some of the membrane-boundenzymes, see Majeran et al. (2008).

Isoprenoids can be synthesized in the cytosolicmevalonate pathway or the chloroplast-localizedMEP pathway (Phillips et al., 2008; Cordoba et al.,2009). Isopentyl diphosphate is an end product of bothpathways and can possibly be transported across thechloroplast envelope. However, it is generally believedthat the MEP pathway provides most, if not all,precursors for the synthesis of carotenoids, chloro-phyll, quinones, and tocopherol. The MEP pathwayand the downstream enzymes geranyl diphosphatesynthase (GPPS) and geranylgeranyl diphosphate syn-thase (GGPS) accumulated at higher levels in the Mthan in BS chloroplasts. This is likely due to the muchhigher demands for carotenoids in the M chloroplasts,as indicated by the low BS-M ratios for several of theenzymes (ZEP, CCD, LUT1, LCY-b; Fig. 9). This higherdemand for carotenoids must relate to their role inquenching of excess light and detoxification of tripletchlorophyll and singlet oxygen, in particular gener-ated during linear electron transport. Similarly, weobserved somewhat preferred M accumulation of theVTE enzymes involved in tocopherol and plastoqui-none synthesis (Fig. 10); both components are neededin the thylakoid membrane, in particular during linearelectron transport.

Nitrogen Assimilation from Inorganic and OrganicNitrogen Sources

Chloroplasts play a central role in nitrogen assimi-lation. During nitrogen assimilation, nitrogen is incor-porated into amino acids in the form of ammonium(Fig. 10; Table III). Plants obtain ammonium from twosources. The primary source originates from inorganicnitrogen, either by reduction of nitrate or by directuptake of ammonium from soil or symbiotic rhizo-bium. The secondary source is derived from organiccompounds within the plant through processes suchas photorespiration. Upon transport of nitrate throughthe vascular system into the BS and M cells, nitrate isreduced to nitrite by nitrate reductase in the cytosol(Lillo, 2008). Nitrite is then imported into the chloro-plast by the nitrite transporter (Sugiura et al., 2007),followed by further reduction into ammonia by Fd-nitrite reductase (NiR) and subsequent incorporationof ammonia into Gln in the GOGAT/GS cycle and itsredistribution to other amino acids through Asptransaminase. An alternative ammonium assimila-tion is catalyzed by the formation of CP for thesynthesis of Arg (Fig. 11) and purimidine nucleotides(Fig. 8).

We did not identify the nitrite transporter in maize,mostly likely due to its low abundance; the Arabidop-sis homolog has not (yet) been identified in any of thepublished proteomics studies either. However, we dididentify and quantify the major nitrogen assimilationenzymes within chloroplasts (Fig. 10; Table III): NiR(BS-M = 0.18), Gln synthase 2 (BS-M = 1.33), Fd-





dependent and NADH-dependent Glu synthases (Fd-GOGAT and NADH-GOGAT; BS-M = 1.00 and 5.33,respectively). Previous studies have indicated thatprimary nitrogen assimilation takes place in M cells,with the Fd-NiR and Fd-GOGAT predominantly local-ized to M cells, while GS activity is present in both celltypes (Rathnam and Edwards, 1976; Harel et al., 1977;Becker et al., 1993). Our proteomic data of NiR and Glnsynthase 2 are consistent with these findings. In the caseof Fd-GOGAT, we found equal abundance in BS and Mcells. The explanation for this apparent discrepancylikely lies in its participation in both primary (prefer-entially in M) and secondary (preferentially in BSfrom photorespiration) nitrogen assimilation. NADH-GOGAT is strongly BS enriched (BS-M = 5.33), but itsoverall abundance is 100-fold lower (based on nSAFvalues) than Fd-GOGAT. NADH-GOGAT is believed tobe generally higher expressed in plant roots than inleaves, but unlike Fd-GOGAT, it has not been studied inmuch detail. The strong preferential cell type-specificaccumulation of NADH-GOGAT likely reflects a met-abolic adaptation to the BS chloroplast environment.

Our results are consistent with the notion that inorganicnitrogen assimilation is tightly correlated with photo-synthesis, and its distribution between BS and M mustdepend on the availability of reducing equivalents forFd-NiR activity.

We detected three related chloroplast envelopetransporters: OMT1 (BS-M = 0.09), DiT1 (BS-M =0.26), and DiT2 (BS-M = 1.16). DiT1 transports 2-oxo-glutarate into the chloroplast, whereas DiT2 exportsGlu into the cytosol; both transporters use malate asthe countertransport molecule (Fig. 10). The preciserole of OMT1 is unclear, but it is clearly preferentiallyaccumulating in M chloroplasts, similar to DiT1.OMT1 and DiT1 are strongly (4- to 10-fold) enrichedin the M chloroplast (BS-M = 0.09 and 0.26, respec-tively), likely because of the high flux rate of malatefrom M chloroplasts in the C4 cycle and low exportfrom BS chloroplasts due to its high rate of malateconsumption by ME. Moreover, photorespiration isconfined to the BS cells (but operates at lower fluxrates than in C3 plants). DiT2 is only slightly higher inBS chloroplasts; using the ZmGI database, we found

Figure 9. BS and M chloroplast distribution of proteins involved in isoprenoid and tetrapyrole metabolism. BS-M bar diagramsand color coding are as described in the legend to Figure 4. Relative accumulation levels (based on nSAF) of proteins averagedacross BS and M chloroplasts are shown as colored squares, with the scale indicated. Abbreviations are explained in the text andin Supplemental Table S3.

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that DiT2 had a higher BS-M ratio (1.7; Majeran et al.,2008; Majeran and vanWijk, 2009). Careful verificationof the full-length sequences of the DiT family isneeded to clarify this difference.Asp aminotransferase catalyzes the reversible trans-

amination between Glu and OAA to give rise to Aspand 2-oxoglutarate. The rate of OAA production in thecytosol (fromPEP by PEP carboxylase) and subsequentimport into M chloroplasts is high, since this is part ofthe C4 cycle. Asp aminotransferase is approximately2-fold higher in M chloroplasts (BS-M = 0.54), consis-tent with our initial observation (Majeran et al., 2005).The production of Asp does compete with productionof malate andmust involve several regulatory steps. Inaddition, Asp aminotransferase may provide a linkbetween nitrogen and carbon pathways in maize, ifAsp is used to transport carbon to the BS cells, as inNAD-ME and PEP carboxykinase types of C4.CPS condenses ammonium (or the Gln amide

group) and HCO32 to make CP, a precursor for Arg

and pyrimidine biosynthesis (Fig. 10). It was indicatedthat, in C3 plants, mitochondrial CPS is involved inrecycling photorespiratory nitrogen (Taira et al., 2004;

Potel et al., 2009). Here, we observed higher levels ofthe CPS large subunit-2 and small subunit in BS cells(BS-M = 4.16 and 2.43 respectively), while the CPSlarge subunit-1 level is comparable (BS-M = 1.03). Thisimplies that the chloroplast-targeted CPS may alsoplay a role in the recovery of photorespiratory nitro-gen. Unlike CPS, most enzymes in the Arg synthesispathway are preferentially located in M cells (seesection on amino acid biosynthesis below); this isbest explained by a lower availability of Glu in BSchloroplasts, as Glu is needed in the GOGAT cycle toremove ammonium released from photorespiration.

In Arabidopsis, nitrogen assimilation is integratedwith carbon metabolism via the chloroplast-localizednitrogen sensor protein PII (Smith et al., 2003), and itinteracts with N-acetyl Glu kinase (Chen et al., 2006).The PII protein senses concentrations of ATP and2-oxoglutarate and relieves Arg feedback inhibitionof N-acetyl Glu kinase. A similar scenario seems inplace in rice (Sugiyama et al., 2004). We identified PIIin Arabidopsis chloroplasts with many MS/MS spec-tra (see PPDB). Surprisingly, there is no obvious maizehomolog of the rice or Arabidopsis PII protein, which

Figure 10. Nitrogen assimilation and Arg metabolism across BS and M chloroplasts. BS-M bar diagrams and color coding are asdescribed in the legend to Figure 4. Relative accumulation levels (based on nSAF) of proteins averaged across BS and Mchloroplasts are shown as colored squares, with the scale indicated. Abbreviations are explained in the text and in SupplementalTable S3.





suggest that integration of carbon and nitrogen me-tabolism is organized differently inmaize and possiblyother C4 species (possibly this integration occursthrough regulating the activity of Asp aminotransfer-ase, as mentioned above).

Sulfur Assimilation and Synthesis of Met, Cys,and Glutathione

Sulfur is absorbed by plants in the form of sulfate,which is first reduced to sulfite and then to sulfidebefore being incorporated into Cys. Sulfate assimila-tion requires high amounts of reducing equivalentsand therefore occurs mostly in the photosyntheticallyactive leaves; reduced sulfur compounds are thendistributed to sink tissues via the phloem (Hopkinset al., 2004; Kopriva and Koprivova, 2005). Sulfatereduction takes place exclusively in chloroplasts,whereas Cys can be synthesized in chloroplasts, cyto-sol, and mitochondria (Krueger et al., 2009, and refs.therein). But since sulfate reduction takes place exclu-sively in plastids in both C3 and C4 species, cytosolic or

mitochondrial Cys synthesis is limited by transport ofsulfide out of the chloroplast (but see below).

We obtained good coverage of enzymes involved inthe step-wise reduction of sulfate to sulfite and then tosulfide (Fig. 11; Table III). Coverage of subsequentsynthesis of Cys, Met, and glutathione was morescattered, likely relating to the distribution of thesesynthetic pathways across multiple subcellular com-partments (Fig. 11). Older studies using other assaysthan proteomics revealed that the activities of ATPsulfurylase (ATPS) and adenosine 5#-phosphosulfatesulfotransferase (APR) in maize were confined to BScells, while Cys synthase (O-acetyl-Ser thiol lyase[OASTL]) activity is found in both cell types but at ahigher level in M cells (Gerwick and Black, 1979;Burnell, 1984; Schmutz and Brunold, 1984; Burgeneret al., 1998). These previous observations are consistentwith our current observations for ATPS2 and APR3 and-4, which showed BS-M ratios of 3.86, 9.92, and 19.75,respectively, whereas twoOASTL isoforms (also namedCys synthase) were 40% to 100% higher in M chloro-plasts than in BS chloroplasts. OASTL was very abun-dant (identified with 504 MS/MS spectra), but we did

Figure 11. Sulfur assimilation and Cys and Met metabolism across BS and M chloroplasts. BS-M bar diagrams and color codingare as described in the legend to Figure 4. Relative accumulation levels (based on nSAF) of proteins averaged across BS and Mchloroplasts are shown as colored squares, with the scale indicated. Abbreviations are explained in the text and in SupplementalTable S3.

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not identify Ser acetyl-CoA transferase (SAT), which isonly active when associated with OASTL. The lack ofdetectable levels of SAT is consistent with a previousreport indicating that OASTL is far more abundant thanSAT (Hopkins et al., 2004). In addition, SAT activity isalso reported to be high in mitochondria and cytosol,and the possibility of OAS transport between subcellu-lar compartments is suggested by experimental data

(Krueger et al., 2009). Interestingly, we found that thehighly abundant sulfite reductase (identified with 172adjSPC) was equally distributed across BS and Mchloroplasts (BS-M = 1.08), suggesting that sulfite istransported from BS to M chloroplasts, in addition towell-established Cys transport (Burgener et al., 1998).Reduction of sulfite into sulfide in M chloroplasts doesdecrease the demand for NADPH in BS, which should

Table III. Distribution of enzymes involved in amino acid metabolism, nitrogen and sulfur assimilation, and iron-sulfur cluster assembly acrossBS and M chloroplasts

Data include annotations for function and subchloroplast location, membrane-stroma ratios (based on nadjSPC) and BS-M ratios (based onnadjSPC), number of matched adjSPC, and relative protein accumulation levels (based on nSAF).

(Table continues on following page.)





be beneficial, as BS chloroplasts lack linear photosyn-thetic electron transport activities (Fig. 11).

Cys is used for the synthesis of Met in a three-stepreaction, involving cystathionine g-synthase, cystathi-onine b-lyase (CBL), and Met synthase (MetS); we didnot observe cystathionine g-synthase, but we foundCBL andMetS preferentially located in BS cells (BS-M =4.72 and 10.01, respectively). CBL clearly appears to bea bona fide chloroplast-localized enzyme (also whencomparing with other maize samples; W. Majeran, G.Friso, and K.J. van Wijk, unpublished data). However,MetS is quite possibly a contamination from the cyto-sol. MetS subcellular localization was believed to becytosolic, but recently it was shown for Arabidopsisthat one of the isoforms of MetS (MS3) is localized inthe plastid, and MetS activity in the plastid wassupported by experimental evidence (Hacham et al.,2008). The MetS protein that we identified was veryabundant in total maize leaf extract and isolated BSstrands, and even if this MetS protein is not with the

chloroplast, it is highly enriched in the BS cells, asindicated in Figure 11.

Cys is also the source for sulfur atoms for proteinswith iron-sulfur clusters; these include several en-zymes in sulfur assimilation (APR and sulfite reduc-tase) as well as enzymes in nitrogen assimilation (NiRand GOGAT) and photosynthetic electron transport(Ye et al., 2006; Xu and Moller, 2008). We identifiedeight proteins involved in iron-sulfur formation: Cyssulfurase (cpSufS/cpNifS) and its activator cpSufEextract elemental sulfur from Cys, cpSufB, -C, and -D,and three NFU proteins that contribute to iron-sulfurcluster assembly (Fig. 11; Table III). These five proteinswere on average 3-fold enriched in the M chloroplast;given the complexity of sulfur metabolism and itsintegration with iron homeostasis, there is no a simpleexplanation for this M enrichment.

The tripeptide glutathione is an important thiolfunctioning in defense and ROS scavenging (see ear-lier section on redox and ROS), and glutathione is also

Table III. (Continued from previous page.)

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a source of electrons for the sulfate reduction (Fig. 11).Glutathione is synthesized from Glu and Gly in a two-step reaction, involving g-glutamyl-Cys synthetase(g-ECS) and glutathione synthase. g-ECS is exclusivelylocalized in plastids in all plant species, whereasglutathione synthase also (or predominantly) occursin the cytosol (as studied in various C3 species). Wedetected g-ECS (with 36 adjSPC) with a larger portionof the total protein located in M chloroplasts (BS-M =0.66), consistent with feeding experiments thatshowed preferential glutathione synthesis in M cells(Burgener et al., 1998). We did not detect glutathionesynthase in the BS or M chloroplasts, which is consis-tent with a report that it is localized in both cytosol andplastids, thus possibly lowering the plastid glutathi-one synthase concentration (Gomez et al., 2004), butwe did detect glutathione reductase at high levels (230adjSPC), responsible for converting oxidized glutathi-one to reduced glutathione. Glutathione reductase was3-fold higher in M chloroplasts, consistent with thehigher rate of ROS production (see section on redoxregulation and ROS defense). Our observations clarifyseveral of the controversies for the distribution ofsulfur metabolism in maize BS and M cells (Koprivaand Koprivova, 2005).

Amino Acid Metabolism

Amino acid synthesis in plants is distributed acrosschloroplasts, cytosol, and mitochondria. Some aminoacids (e.g. Gln) act primarily to assimilate and trans-port nitrogen. Amino acids are also precursors ofmany nitrogen-containing compounds. Therefore,amino acid biosynthesis is regulated through variousmechanisms and with various alternative pathwaysand is tightly coordinated with carbon metabolism.Given this complexity, it is perhaps not surprising thatvery little is known about the distribution of aminoacid biosynthetic pathways between BS and M cells inC4 plants. In our previous BS-M stromal analysis, wecould only identify a handful of enzymes involved inamino acid biosynthesis (Majeran et al., 2005). In thisstudy, we identified and functionally assigned some 60proteins/protein groups (about 70 genes) involved inamino acid metabolism (with an overlap to sulfur andnitrogen assimilation and photorespiration), and theycovered 3 orders of magnitude in relative concentra-tion (based on nSAF values; Table III). Amino acidbiosynthetic pathways can be separated based on theprecursor, but connections between these pathwaysexist. We have grouped the identified proteins indifferent pathways in Table III, and a number ofproteins and pathways are integrated in the figuresfor nitrogen and sulfur assimilation (e.g. for Arg andCys biosynthesis).Asp aminotransferase was the most abundant pro-

tein (based on nSAF; 1,687 MS/MS spectra; BS-M =0.54), followed by shikimate kinase SKL1 (488 MS/MSspectra; BS-M = 0.7) involved in biosynthesis of aro-matic amino acids. Among the proteins with at least 40

adjSPC, we observed BS-M ratios ranging from 0.06(greater than 10-fold enriched in M chloroplasts) to5.96 (6-fold enriched in BS chloroplasts), with themajority of proteins being enriched in M chloroplasts(Table III). In the remainder of this section, we willbriefly describe our findings, with an emphasis on thelink to C4 metabolism.

For the Glu family (Glu, Arg, and Pro), we identifiedseven out of the eight enzymes needed for Arg bio-synthesis. Arg is synthesized from Glu and CP and isthus closely linked to the GOGAT/GS cycle. All sevenidentified Arg biosynthetic enzymes were enriched inthe M chloroplasts (on average 2.5-fold; Table III; Fig.10). There must be competition between Arg andnucleotide synthesis, as both require CP as a precursor.We suggest that Arg synthesis and accumulation serveto redistribute excess assimilated nitrogen to otherparts of the plant. It is interesting that in the Arabi-dopsis glu1 mutant, excess photorespiratory ammo-nium was transported in the form of Arg (and otheramino acids; Potel et al., 2009).

Asp is the precursor for homoserine, Thr, Ile, andLys and also contributes to the synthesis of Met. Weidentified nine enzymes in these pathways, with theLys synthesis branch being well covered (Table III).The pathway was M enriched, which is understand-able given the high rates of Asp synthesis and highenergy requirements (ATP and NADPH).

We identified most of the enzymes involved insynthesis of the branched amino acids, Val and Leu(Table III). The precursor of this pathway is pyruvate,and the last step in their synthesis requires Glu as anamine source. The later steps in Leu synthesis areclearly not M enriched but either equally distributedacross both cell types or enriched in the BS cell; thereason is not clear.

As shown in Figure 11, we identified several of theenzymes involved in Ser and Cys synthesis (Table III),as explained in the section on sulfur assimilation. Wedid not identify proteins involved in Gly synthesis,which is reassuring, since Gly is primarily produced inmitochondria from Thr, Ser, or photorespiration.

The shikimate pathway for the synthesis of thearomatic residues (Phe, Tyr, and Trp) was well coveredwith 11 proteins, even if the number of SPC was lowfor several of the steps (see “Conclusion”; Table III).The entire shikimate pathway takes place in plastidsand starts with the condensation of erythrose 4-phos-phate (a product of the pentose phosphate pathwayand of the Calvin cycle) and PEP, followed by incor-poration of a second molecule of PEP in the penulti-mate step of the central pathway. This central part ofthe pathway produces chorismate in seven steps,followed by two separate pathways for the synthesisof Trp and of Tyr plus Phe. We identified three steps (1,5, and 6) in the central pathway and multiple steps inthe Trp- and Tyr/Phe-specific pathways. All but one ofthe enzymes were more highly expressed in the Mchloroplasts (the BS-enriched exception was identifiedby only four SPC and therefore is not reliable). EPSP





synthase (for 3-phosphoshikimate 1-carboxyvinyl-transferase), in the central part of the shikimate path-way, is the target of the well-known and extremely“popular” herbicide glyphosate; EPSP synthase was5-fold enriched in M chloroplasts. The preferentialaccumulation in M chloroplasts is in line with the highlevels of PEP synthesis in the M chloroplasts for the C4carbon cycle. The availability and origin of erythrose4-phosphate in the M chloroplasts is less clear butmust be generated by the pentose phosphate pathwayrather than the Calvin cycle. Indeed, this is consistentwith the observed accumulation of OPPP enzymes inthe M chloroplasts (G6PDH, lactonate, and 6PGDH;Fig. 5). The last step in Trp synthesis is carried out bysubunits of the Trp synthase complex; these subunitswere 3- to 4-fold higher in M than in BS chloroplasts(Table III).

The synthesis of His follows a linear pathway,originating with phosphoribosyl pyrophosphate (Fig.8). We identified enzymes for six out of eight steps ofthe pathway, even if the number of SPC was low forall but the last step, histidinol dehydrogenase (59matchedMS/MS spectra). The BS-M ratio of histidinoldehydrogenase was 0.66, but the distribution of thecomplete His pathway across both cell types is notclear. Clearly, the His pathway in C4 species requiresmore attention.

Many Proteins of Unknown Function Show Differential

BS-M Accumulation

In addition to the proteins discussed above, weidentified and quantified over 200 proteins (and groups)with either a miscellaneous function (e.g. TPR andPPR proteins, rhodanese, and DnaJ domain proteins)or without any obvious function (Supplemental TableS3). Ninety-six proteins were only detected in the stro-mal fractions, and 25 were identified in both stromaand membrane fractions. These proteins spanned 3orders of abundance (based on nSAF) and up to 1,556matched spectra; a significant number of proteinsshowed strong preferential BS or M accumulationand are an excellent resource for further explorationof specialized BS or M chloroplast functions.

CONCLUSION

Large-scale MS analysis with a high-resolution andhigh-sensitivity mass spectrometer has allowed path-ways and activities to be analyzed and reconstructedacross the differentiated BS and M chloroplasts in theleaf of maize. Quantitative information about relativeprotein accumulation levels and cell type-specificprotein accumulation patterns provided new insightsinto the functions and structures of differentiated BSand M chloroplasts. These results provide a majorstep forward from previous analyses and also givemany new entry points for studying specific aspectsof chloroplast biogenesis, differentiation, and func-tion. We did note that a number of proteins, partic-

ularly in the functional categories of nucleotidemetabolism and DNA interactions and transcription,were quantified with only a few spectra counts. Thisis likely due to a down-regulation of these functionswhen chloroplasts begin to reach the end point ofbiogenesis and differentiation. For instance, nonpho-tosynthetic chloroplasts (etioplasts) at the leaf basewill be expected to contain much higher proteinlevels for these functions.

The availability of the first maize genome sequencehas provided an excellent template for protein identi-fication and quantification, even if imperfectionssurely must have resulted in missed proteins andinaccuracies of protein quantification. Through thisstudy, we have added a wealth of information to thesemaize protein accession numbers, and this includesassignment of protein names, functions, and subcellu-lar localizations; this will all be made available onlinethrough the PPDB. From PPDB, this information canbe 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 thepublic depository Proteomics Identifications (PRIDE;http://www.ebi.ac.uk/pride/).

A central conclusion from this study is that differ-entiated BS chloroplasts at the maize leaf tip appearto have strongly reduced plastid protein expressionand protein import. Moreover, it is likely activelyremodeling its proteome, as evidenced by increasedlevels of ClpB and HSP90. Furthermore, this studyprovides strong experimental support for a numberof specialized BS and M metabolic functions, includ-ing starch biosynthesis (BS), Arg synthesis (M), MEPactivity (M), nitrogen assimilation (M), initial steps insulfur assimilation (BS), and more. Functional andgenetic studies, as well as in vitro enzyme activityassays, will be needed to further determine theirsignificance. In a forthcoming study, we will explorethe protein accumulation of the various pathwaysand processes along the leaf developmental gradientand strengthen and expand on our observationspresented here for the differentiated BS and M chlo-roplasts.

MATERIALS AND METHODS

Maize Genotype, Plant Growth, and Purification of BSand M Chloroplast Fractions

WT-T43 maize (Zea mays) plants were grown for 12 to 14 d in a growth

chamber (16 h of light/8 h of dark, 400 mmol photons m22 s21) until the fourth

leaf was emerging. M and BS chloroplasts were purified from the top 4-cm

section of the third leaf and harvested about 2 h after the onset of the light

period, using several hundred leaf tips, following procedures described

previously (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 of centrifugation at 80,000g. The supernatant, representing

the enriched soluble stromal cross-contamination of M and BS chloroplast

fractions, was collected. Cross-contamination was assessed from the presence

of the M and BS markers (PPDK and Rubisco, respectively) as visualized on

Friso et al.




stained one-dimensional SDS-PAGE gels, as described (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 Analysisby NanoLC-LTQ-Orbitrap

A total of 150 mg of each BS and M stromal preparation was separated by

SDS-Tricine-PAGE (12% acrylamide). Each gel lane was then cut into 12 slices,

proteins were digested with trypsin, and the extracted peptides were ana-

lyzed by nanoLC-LTQ-Orbitrap MS using data-dependent acquisition and

dynamic exclusion, as described (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 carryover of peptides that could

bias quantification. Purification and MS analysis of the BS and M membranes

has been described (Majeran et al., 2008).

Processing of the MS Data, Database Searches, andUpload into PPDB

Peak lists (.mgf format) were generated using DTA supercharge (version

1.19) software (http://msquant.sourceforge.net/) and searched with Mascot

version 2.2 (Matrix Science) against maize genome release 4a.53 (with 53,764

models) from http://www.maizesequence.org/ supplemented with the plastid-

encoded proteins (111 protein models) and mitochondria-encoded proteins

(165 protein models). For off-line calibration, first a preliminary search was

conducted with the precursor tolerance window set at 630 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 then applied to adjust

precursor masses in the peak lists of the respective .mgf file for recalibration

using a Perl script (B. Zybailov, unpublished data). The recalibrated peak lists

were searched against the maize genome data set with organellar genes (with

54,380 entries) and in parallel against ZmGI version 16.0 (with 56,364 entries),

including sequences for known contaminants (e.g. keratin, trypsin) concate-

nated with a decoy database where all the sequences were randomized. Each

of the peak lists were searched using Mascot version 2.2 (maximum P of 0.01)

for full tryptic peptides using a precursor ion tolerance window set at 66

ppm, variable Met oxidation, fixed Cys carbamidomethylation, and a minimal

ion score threshold of 30 for maize genome and 44 for ZmGI; this yielded a

peptide false discovery rate below 1%, with peptide false positive rate

calculated as: 2 3 (decoy_hits)/total_hits. The false protein identification

rate of protein identified with two or more peptides was zero. To reduce the

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 63 ppm. Precursor ion masses below

700 D were discarded. All filtered results were uploaded into 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 PRIDE database at

http://www.ebi.ac.uk/pride/.

Selection of the Best Gene Models and Post-Mascot Filterto Assign Shared and Unique Peptides, and Creation ofProtein Groups with a High Percentage of Shared

Matched Spectra

Many genes have more than one 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 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 SPC for each accession as a basis.

Proteins that shared more than approximately 80% of their matched adjusted

peptides with other proteins across the complete data set 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 \ 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 (e.g. less than 20). The linkage of homologous proteins that were

identified by the same set of MS/MS spectra was recorded (their identification

was marked as “ambiguous”), and the matched MS/MS spectra were marked

as “shared” spectra (or “not unique”). For proteins that were identified 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 Protein BS-MRatios Using nSAF Values

Relative abundance for each identified protein accession (or cluster) was

calculated by the 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 to 3,500 D, since shorter peptides were excluded

from the Mascot search results, while the longer peptides were beyond the

mass-to-charge ratio 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 membrane-

enriched fractions as compared with 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 (Supplemental

Tables S2 and S3).

The PPDB and Functional Assignment ofIdentified Proteins

MS-based information of all identified proteins was extracted from the

Mascot search pages and filtered for significance (e.g. minimum ion scores,

etc.), ambiguities, and shared spectra as described (Zybailov et al., 2008). This

information includes Mowse scores, number of matching peptides, number of

matched MS/MS spectra (counts), number of unique and adjusted counts,

highest peptide score, highest peptide error (ppm), lowest absolute error

(ppm), sequence coverage, and tryptic peptide sequences. This information is

available in the PPDB (Sun et al., 2009) 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”; 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 names of ZmGI and

maize genome protein accessions was based on a combination of best BLAST

hits in the predicted rice (Oryza sativa) proteome (OsGI version 5; from http://

rice.plantbiology.msu.edu/), the predicted Arabidopsis (Arabidopsis thaliana)

proteome (ATH version 8 from The Arabidopsis Information Resource

[http://www.arabidopsis.org/]), and our manual annotations for ZmGI

version 16 (from http://compbio.dfci.harvard.edu/) homologues (Majeran

et al., 2005, 2008; Majeran and van Wijk, 2009). Pair-wise BLAST search results

between ATH version 8, OSGI version 5, and ZmGI version 16 are available

via PPDB. Each identified protein was assigned to a molecular function using

the hierarchical, nonredundant classification system developed for MapMan

(Thimm et al., 2004; http://gabi.rzpd.de/projects/MapMan/), adjusted after

manual verification and information from the literature, and incorporated into

PPDB. Predicted PFAM domains for all predicted maize genome proteins are

also available in PPDB.

Supplemental Data

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





Supplemental Figure S1. BS-M protein accumulation ratios of proteins of

the thylakoid-associated photosynthetic apparatus.

Supplemental Table S1. Identification of 2,439 protein models in stroma-

enriched and membrane-enriched fractions from BS and M chloro-

plasts.

Supplemental Table S2. Identification, annotation, and filtering of the

1,429 best gene models.

Supplemental Table S3. Final data set with 923 proteins and protein

groups with final annotations, nSAF and nadjSPC values, BS-M ratios,

membrane-stroma ratios, the matched amino acid sequences, and

Arabidopsis and rice best homologues.

Supplemental Text S1. Detailed explanation of Table I.

Received December 23, 2009; accepted January 17, 2010; published January 20,

2010.

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