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Proteome Analysis of the Rice Etioplast

2005; Elsevier BV; Volume: 4; Issue: 8 Linguagem: Inglês

10.1074/mcp.m500018-mcp200

1535-9484

Anne von Zychlinski, Torsten Kleffmann, Nandini Krishnamurthy, Kimmen Sjölander, Sacha Baginsky, Wilhelm Gruissem,

Microbial Metabolic Engineering and Bioproduction

We report an extensive proteome analysis of rice etioplasts, which were highly purified from dark-grown leaves by a novel protocol using Nycodenz density gradient centrifugation. Comparative protein profiling of different cell compartments from leaf tissue demonstrated the purity of the etioplast preparation by the absence of diagnostic marker proteins of other cell compartments. Systematic analysis of the etioplast proteome identified 240 unique proteins that provide new insights into heterotrophic plant metabolism and control of gene expression. They include several new proteins that were not previously known to localize to plastids. The etioplast proteins were compared with proteomes from Arabidopsis chloroplasts and plastid from tobacco Bright Yellow 2 cells. Together with computational structure analyses of proteins without functional annotations, this comparative proteome analysis revealed novel etioplast-specific proteins. These include components of the plastid gene expression machinery such as two RNA helicases, an RNase II-like hydrolytic exonuclease, and a site 2 protease-like metalloprotease all of which were not known previously to localize to the plastid and are indicative for so far unknown regulatory mechanisms of plastid gene expression. All etioplast protein identifications and related data were integrated into a data base that is freely available upon request. We report an extensive proteome analysis of rice etioplasts, which were highly purified from dark-grown leaves by a novel protocol using Nycodenz density gradient centrifugation. Comparative protein profiling of different cell compartments from leaf tissue demonstrated the purity of the etioplast preparation by the absence of diagnostic marker proteins of other cell compartments. Systematic analysis of the etioplast proteome identified 240 unique proteins that provide new insights into heterotrophic plant metabolism and control of gene expression. They include several new proteins that were not previously known to localize to plastids. The etioplast proteins were compared with proteomes from Arabidopsis chloroplasts and plastid from tobacco Bright Yellow 2 cells. Together with computational structure analyses of proteins without functional annotations, this comparative proteome analysis revealed novel etioplast-specific proteins. These include components of the plastid gene expression machinery such as two RNA helicases, an RNase II-like hydrolytic exonuclease, and a site 2 protease-like metalloprotease all of which were not known previously to localize to the plastid and are indicative for so far unknown regulatory mechanisms of plastid gene expression. All etioplast protein identifications and related data were integrated into a data base that is freely available upon request. Plastids are plant cell organelles that have essential biosynthetic and metabolic activities. These include photosynthetic carbon fixation and synthesis of amino acids, fatty acids, starch, and secondary metabolites such as pigments. Although plastids lost their autonomy and transferred most of their genes to the nucleus during evolution (1Martin W. Herrmann R.G. Gene transfer from organelles to the nucleus: how much, what happens, and why?.Plant Physiol. 1998; 118: 9-17Google Scholar), they have retained a small genome encoding ∼90 proteins. Different plastid types develop in a tissue-specific manner (for a detailed review on plastid biogenesis, see Ref. 2Vothknecht U.C. Westhoff P. Biogenesis and origin of thylakoid membranes.Biochim. Biophys. Acta. 2001; 1541: 91-101Google Scholar). According to their structure, pigment composition (color), and functional differentiation, plastids are classified as elaioplasts that are found in seed endosperm, chromoplasts in fruits and petals, amyloplasts in roots, etioplasts in dark-grown seedlings, and chloroplasts in photosynthetically active tissues (3Neuhaus H.E. Emes M.J. Nonphotosynthetic metabolism in plastids.Annu. Rev. Plant Physiol. Plant Mol. Biol. 2000; 51: 111-140Google Scholar). These specialized plastids types are typically the result of a differentiation program that is controlled by the cell and tissue type but also by environmental factors. Perhaps the best understood example of plastid differentiation is the light-dependent conversion of etioplasts into chloroplasts. After exposure of dark-grown seedlings to light etioplasts differentiate into photosynthetically active chloroplasts within a few hours. Chloroplast differentiation is accompanied by the assembly of the thylakoid membrane-localized electron transport system, which requires proteins encoded by genes in both nuclear and chloroplast genomes (4Gray J.C. Sullivan J.A. Wang J.H. Jerome C.A. MacLean D. Coordination of plastid and nuclear gene expression.Philos. Trans. R. Soc. Lond. B Biol. Sci. 2003; 358: 135-145Google Scholar, 5Jarvis P. Intracellular signalling: the language of the chloroplast.Curr. Biol. 2003; 13: R314-R316Google Scholar). Although chloroplast differentiation has been investigated in detail for many years, the molecular mechanisms that control the differentiation are not yet fully understood. Also information is still limited on the metabolic pathways that distinguish plastids in different heterotrophic tissues and from photosynthetically active chloroplasts. Using proteome analysis we examined the global state of protein expression in etioplasts to establish comprehensive information on complex metabolic and regulatory networks that function in a heterotrophic plastid. Proteome analysis has become an indispensable source of information about protein expression, splice variants, and erroneous or incomplete prediction of gene structures in data bases. The analyses of cell organelle proteomes provide additional important information about protein localization and pathway compartmentalization. Most of the currently available plastid proteome information that provides new insights into organelle-specific metabolic functions has been reported from autotrophic chloroplasts (6Peltier J.B. Friso G. Kalume D.E. Roepstorff P. Nilsson F. Adamska I. van Wijk K.J. Proteomics of the chloroplast: systematic identification and targeting analysis of lumenal and peripheral thylakoid proteins.Plant Cell. 2000; 12: 319-341Google Scholar, 7Ferro M. Salvi D. Riviere-Rolland H. Vermat T. Seigneurin-Berny D. Grunwald D. Garin J. Joyard J. Rolland N. Integral membrane proteins of the chloroplast envelope: identification and subcellular localization of new transporters.Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 11487-11492Google Scholar, 8Peltier J.B. Emanuelsson O. Kalume D.E. Ytterberg J. Friso G. Rudella A. Liberles D.A. Soderberg L. Roepstorff P. von Heijne G. van Wijk K.J. Central functions of the lumenal and peripheral thylakoid proteome of Arabidopsis determined by experimentation and genome-wide prediction.Plant Cell. 2002; 14: 211-236Google Scholar, 9Schubert M. Petersson U.A. Haas B.J. Funk C. Schroder W.P. Kieselbach T. Proteome map of the chloroplast lumen of Arabidopsis thaliana.J. Biol. Chem. 2002; 277: 8354-8365Google Scholar, 10Ferro M. Salvi D. Brugiere S. Miras S. Kowalski S. Louwagie M. Garin J. Joyard J. Rolland N. Proteomics of the chloroplast envelope membranes from Arabidopsis thaliana.Mol. Cell. Proteomics. 2003; 2: 325-345Google Scholar, 11Froehlich J.E. Wilkerson C.G. Ray W.K. McAndrew R.S. Osteryoung K.W. Gage D.A. Phinney B.S. Proteomic study of the Arabidopsis thaliana chloroplastic envelope membrane utilizing alternatives to traditional two-dimensional electrophoresis.J. Proteome Res. 2003; 2: 413-425Google Scholar, 12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar, 13Lonosky P.M. Zhang X. Honavar V.G. Dobbs D.L. Fu A. Rodermel S.R. A proteomic analysis of maize chloroplast biogenesis.Plant Physiol. 2004; 134: 560-574Google Scholar, 14Peltier J.B. Ytterberg A.J. Sun Q. van Wijk K.J. New functions of the thylakoid membrane proteome of Arabidopsis thaliana revealed by a simple, fast, and versatile fractionation strategy.J. Biol. Chem. 2004; 279: 49367-49383Google Scholar). Here we report a novel protocol for the isolation of highly purified rice etioplasts and subsequent systematic analysis of the etioplast proteome. The proteins we identified reveal a complex etioplast-specific metabolism and novel regulatory functions in plastids during heterotrophic growth and early differentiation of the autotrophic chloroplast. Our results represent an important first step toward an integrated view of plastid metabolism and differentiation processes. For each plastid isolation 200 g of rice (Oryza sativa, japonica cultivar group) seeds were washed in 5% sodium hydrochloride solution for 10 min, rinsed four times with water, and swollen overnight. The seeds were transferred to vermiculite watered with half-concentrated Murashige and Skoog medium. Seedlings were grown in the dark at a constant 29 °C for 10 days. Plastids were purified by a combination of two consecutive density gradient centrifugations using Nycodenz (Axis-Shield PoC AS, Oslo, Norway) as the density gradient medium and several differential centrifugation steps. Each step of the isolation procedure was carried out at 4 °C. Rice shoots were harvested and cut into 5–10-mm lengths. In batches of 50 g, the plant material was homogenized in 500 ml of etioplast isolation solution (E.I.S.) 1The abbreviations used are: E.I.S., etioplast isolation solution; 2-D, two-dimensional; MIP, major intrinsic protein; BY-2, Bright Yellow 2; OPPP, oxidative pentose phosphate pathway; PPT, phosphoenolpyruvate/phosphate translocator; TPT, triose phosphate/phosphate translocator; PEP, phosphoenolpyruvate; POR (A and B), protochlorophyllide oxidoreductase (isoforms A and B); Tic, translocon of the inner envelope membrane of chloroplasts; Toc, translocon of the outer envelope membrane of chloroplasts; Tim, translocon of the inner membrane of mitochondria; CSP41, 41-kDa chloroplast stem-loop-binding protein; S2P, site 2 protease; SREBP, sterol regulatory element-binding protein; E-value, Expect value. containing 10 mm HEPES/KOH, pH 7.8, 2 mm EDTA, 2 mm MgCl2, 1 mm tetrasodium pyrophosphate, 600 mm sorbitol, 0.2% (w/v) BSA (Fraction V, Sigma) with a Waring blender (three consecutive bursts at low speed and three at high speed). The homogenate was filtered through two layers of Miracloth (Calbiochem). The homogenization step was repeated once. The pooled filtrates were refiltered through four layers of Miracloth. The filtrate was subsequently centrifuged for 4 min at 200 × g to remove cellular debris. The supernatant was recentrifuged for 10 min at 8000 × g. All centrifugation steps were carried out at 4 °C with a Sorvall RC5C centrifuge (DuPont). The pellets containing the plastids were carefully resuspended in E.I.S. and subjected to Nycodenz density gradient centrifugation. The supernatant of the differential centrifugation was centrifuged at 16,000 × g for 20 min, and the resulting pellet containing a mixture of various cell organelles was put aside for further analysis. A 50% (w/v) Nycodenz stock solution (containing 10 mm HEPES/KOH, pH 7.8, 2 mm EDTA, 2 mm MgCl2, 1 mm tetrasodium pyrophosphate, 5 mm DTT) was diluted with E.I.S. (plus 5 mm DTT) to the required Nycodenz concentration and an osmolality of 500–550 mosm. The plastid suspension was adjusted with 50% Nycodenz stock solution to a final Nycodenz concentration of 30%. Five milliliters of the suspension were loaded in a 30-ml Corex tube (Number 8445). For 200 g of rice seeds we used four Corex tubes for the first Nycodenz gradient. The Nycodenz step gradient consisting of 6 ml of 25%, 8 ml of 20%, 6 ml of 15%, and 3 ml of 10% Nycodenz was then carefully casted on top of the organelle suspension. The gradient was centrifuged in a Sorvall HB-4 swinging bucket rotor at 8000 × g for 45 min. Two yellowish bands (bands 2 and 3) at the interface of 20–15 and 25–20% Nycodenz and four pale grayish bands (bands 0, 1, 4, and 5) at the interface of 0–10, 10–15, and 25–30% and in the pellet of slightly higher than 30% Nycodenz were visible. All bands were examined by light microscopy using an Axioplan 2 microscope (Zeiss, Wetzikon, Switzerland). Bands 2 and 3 contained the highest amount of plastids and were used for further purifications. Bands 0, 1, 4, and 5 were diluted 3-fold with E.I.S. and centrifuged at 16,000 × g for 20 min. The resulting pellets containing a mixture of various organelles were stored for additional analysis. Bands 2 and 3 were pooled and diluted 3-fold (v/v) with E.I.S. plus 5 mm DTT and subjected to differential centrifugation carried out in an SS34 rotor as described below. 1) The organelle suspension was centrifuged for 5 min at 8000 × g to remove residual Nycodenz. The resulting pellet was resuspended in a maximum of 20 ml of E.I.S. per centrifuge tube and used for the second centrifugation step. 2) The second centrifugation was carried out for 10 min at 500 × g. The resulting pellet was resuspended as before and used for the third centrifugation step. 3) The third centrifugation was carried out for 15 min at 500 × g, resulting in the first plastid pellet. To increase the yield the supernatant from the second centrifugation was recentrifuged for 10 min at 500 × g. The resulting pellet was resuspended, and the centrifugation step was repeated for 15 min at 500 × g, resulting in the second plastid pellet. Both pellets were combined and subjected to the second Nycodenz gradient centrifugation using the same composition as described for the first gradient (see above). Due to the low amount of resulting material we used only one centrifuge tube for the second gradient. After the second gradient bands 2 and 3 were collected separately, diluted 3-fold (v/v) with E.I.S., and centrifuged for 5 min at 8000 × g to remove residual Nycodenz. The pellets were resuspended in a maximum of 20 ml of E.I.S. and centrifuged using an SS34 rotor for 10 min at 2000 × g. The final pellets were stored at −80 °C. The pellets from the organelle fractionation, i.e. the pellet from band 2 of the second density gradient (plastid fraction) as well as the pellets from the supernatant of the first centrifugation step and from bands 0, 1, 4, and 5 after the first density gradient ("unclean" fractions) were resuspended in SDS sample buffer (0.675 mm Tris/HCl, pH 6.8, 10% glycerol, 20% 2-mercaptoethanol, bromphenol blue) and incubated at 30 °C for 30 min. Any insoluble material was pelleted for 10 min at 16,000 × g. Two hundred micrograms of solubilized proteins were directly subjected to SDS-PAGE by loading 50 μg/lane onto 10-cm-long homogeneous 12% polyacrylamide gels. After electrophoresis the gels were cut into 10 segments along the lane. Proteins in each gel segment were immediately subjected to in-gel tryptic digest as described previously (15Shevchenko A. Wilm M. Vorm O. Mann M. Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels.Anal. Chem. 1996; 68: 850-858Google Scholar) using sequencing grade modified trypsin (Promega) in a ratio of one part trypsin to 10 parts of protein. Proteins were digested overnight at 26 °C. After elution, tryptic peptides were lyophilized to dryness and stored at −80 °C until analysis. Tryptic peptides of each fraction were resuspended in 5 μl of 5% ACN and 0.2% formic acid in water and loaded on laboratory-made silica capillary columns (inner diameter of 75 μm, length of 9 cm; BGB Analytik AG, Böckten, Switzerland) packed with C18 reversed phase material (Magic C18 resins; 5 μm, 200-Å pore; Michrom BioResources, Auburn, CA). The peptide mixture was separated and eluted by a gradient from 5 to 65% ACN over 2 h followed by an increase up to 80% during an additional 15 min. The flow rate at the tip of the column was adjusted to ∼200 nl/min. LC was coupled on line to an LCQDeca XP ion trap mass spectrometer (Thermo Finnigan, San Jose, CA) equipped with a nanospray source. Mass analysis was performed with a spray voltage of 2.0–2.5 kV and one MS full scan followed by three data-dependent MS/MS scans of the three most intense parent ions. The dynamic exclusion function was enabled to allow two measurements of the same parent ion during 1 min followed by an exclusion duration of 1 min. MS/MS data were interpreted according to the standards put forward by Carr and colleagues (16Carr S. Aebersold R. Baldwin M. Burlingame A. Clauser K. Nesvizhskii A. The need for guidelines in publication of peptide and protein identification data: Working Group on Publication Guidelines for Peptide and Protein Identification Data.Mol. Cell. Proteomics. 2004; 3: 531-533Google Scholar). The SEQUEST software was used (Thermo Finnigan) to search the O. sativa protein data base (TIGR: The Institute of Genome Research, www.tigr.org/) including the plastid- and mitochondria-encoded proteins (download from May 5, 2004). dta files were created by the SEQUEST software for every MS/MS scan with a total ion count of at least 5 × 104, minimal peak count of 35, and a precursor ion mass in the range of 300–2000 m/z. Data were searched against the data base indexed for speed restricting the search to tryptic peptides without modifications (carboxyamidomethylated cysteines and oxidized methionines). To exclude any false positive protein identification we manually interpreted each SEQUEST output by filtering the peptide hits using stringent hierarchical criteria as described previously (12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar). In brief, we accepted cross-correlation scores (Xcorr) of at least 2.5 for doubly and 3.5 for triply charged ions. Grouping of at least four significant peptide hits of the same protein was rated as significant protein identification. MS/MS spectra of the other protein hits were visually examined for a correct peak assignment considering criteria described previously (12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar). For peptide identifications with a ΔCN (normalized difference in correlation score, giving the differences between the front ranking and the following possible hit) lower than 0.1, the spectra of lower ranking hits were also examined. Identifications with a ΔCN of 0.0 resulting from different members of protein families, isoforms, or redundant data base entries that cannot be distinguished by the identified peptides are given in Supplemental Table 1B. In one case where we based a biological conclusion on a single hit protein identification, we searched the dta file with MASCOT (www.matrixscience.com) using the default parameters (error tolerance of 2 for the parent mass and 0.8 for the daughter ions) to confirm the SEQUEST identification. This MS/MS spectrum was identified with a MASCOT score of 78 as a peptide from the site 2 metalloprotease supporting the SEQUEST result (Supplemental Fig. 1). For 2-D PAGE plastid pellets of bands 2 and 3 were resuspended each in approximately 100 μl of solubilization buffer containing 40 mm Tris base, 7 m urea, 2 m thiourea, 2% CHAPS, 0.5% Brij 35, 0.4% carrier ampholytes, 2 mm tributyl phosphine (Fluka, Buchs, Switzerland), 20 mm DTT, complete EDTA-free protease inhibitor mixture (Roche Applied Science) resulting in a protein concentration of at least 1 μg/μl. Any insoluble material was pelleted for 30 min at 20,000 × g. For the first dimension 100 μg of protein were loaded onto 24-cm-long strips with an immobilized linear pH gradient from 4–7 (Bio-Rad) by in-gel rehydration. The rehydration was performed overnight in solubilization buffer without Tris base according to the manufacturer's instructions. Proteins were focused using the IPGphor (Amersham Biosciences) by the following voltage gradient: 2 h at 300 V, 1 h up to 600 V, 1 h up to 1000 V, 1 h up to 4000 V, and for a total of approximately 80,000 V-h at 4000 V. Focused strips were stored at −80 °C. The second dimension was performed in laboratory-made homogeneous 12% polyacrylamide gels using the Ettan Dalt 2 unit (Amersham Biosciences). Proteins were detected by SYPRO® Ruby staining (Molecular Probes Europe BV, Leiden, The Netherlands) and scanned with a Typhoon 9400 scanner (Amersham Biosciences). 2-D PAGE electropherograms were analyzed and compared with the ProteomWeaver software (Definiens, Munich, Germany). All relevant data of the 240 identified etioplast proteins were deposited in a relational data base designated as PLprot. This data base is accessible at www.pb.ethz.ch/proteomics and also contains information about the Arabidopsis chloroplast proteome as described previously (12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar). The TargetP program (www.cbs.dtu.dk/services/TargetP/) was used to predict subcellular protein targeting for plastids, mitochondria, the secretory pathway, and any other localization. For the identification of protein homologies the BLAST searches were conducted as described previously (17Altschul S.F. Madden T.L. Schaffer A.A. Zhang J. Zhang Z. Miller W. Lipman D.J. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs.Nucleic Acids Res. 1997; 25: 3389-3402Google Scholar) against either the NCBI non-redundant data base (www.ncbi.nlm.nih.gov/BLAST/) or the Swiss-Prot data base using the BLAST program (bio.thep.lu.se/). Domain and structure homologies were analyzed either by the conserved domain search at NCBI (www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) using the Conserved Domain Database version 2.00 or with phylofacts (phylogenomics.berkeley.edu/resources/). To provide more informative annotations of proteins without annotated function we further attempted informatic approaches as described previously (12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar). Briefly we performed a four-step procedure with the first two steps being essentially identical to those used in the functional classification of the human genome (18Venter J.C. Adams M.D. Myers E.W. Li P.W. Mural R.J. Sutton G.G. Smith H.O. Yandell M. Evans C.A. Holt R.A. et al.The sequence of the human genome.Science. 2001; 291: 1304-1351Google Scholar). The third step focused on the prediction of structural domains in the proteins using fold recognition methods for the critical assessment of protein structure prediction (CASP) experiments (19Karplus K. Sjolander K. Barrett C. Cline M. Haussler D. Hughey R. Holm L. Sander C. Predicting protein structure using hidden Markov models.Proteins. 1997; : 134-139Google Scholar). In the fourth step we finally used publicly available software tools (www.tigr.org/, www.ncbi.nlm.nih.gov/, www.cbs.dtu.dk/services/TargetP/, www.sanger.ac.uk/Software/Pfam/index.shtml, phylogenomics.berkeley.edu/resources, scop.berkeley.edu, www.rcsb.org/pdb, bioweb.pasteur.fr/seqanal/interfaces/toppred.html, bio.thep.lu.se/, and bibiserv.techfak.uni-bielefeld.de/dialign/) to complement our structure and function prediction analyses. A detailed description of the complete procedure was recently published for the annotation of protein functions for Arabidopsis chloroplast proteins (12Kleffmann T. Russenberger D. von Zychlinski A. Christopher W. Sjolander K. Gruissem W. Baginsky S. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions.Curr. Biol. 2004; 14: 354-362Google Scholar). Large scale analysis of the etioplast proteome has been constrained by the lack of suitable strategies to isolate highly purified etioplasts from dark-grown plant tissue. This is of particular concern because protein detection has become more sensitive. Minor contaminations with proteins from other cell organelles might therefore result in misinterpretation of the proteome data. To reduce ambiguities in the assignment of proteins to the etioplast proteome, we first tested different established plastid isolation methods for the purification of rice etioplasts. Sucrose density gradient centrifugation yielded plastids contaminated with other cell organelles and particles (data not shown) and exposed plastids to strong osmotic stress, which in general makes the isolation of intact plastids difficult. Percoll density gradient centrifugation resulted in the aggregation of rice etioplasts, which prevented their efficient separation from other cell organelles (data not shown). In contrast, a novel protocol that combines differential centrifugation and Nycodenz density gradient centrifugation (for details see "Experimental Procedures") allowed the isolation of intact and pure etioplasts from leaves of dark-grown rice seedlings. After two consecutive Nycodenz density gradient centrifugations of leaf extracts two light yellow bands (bands 2 and 3) were visible at the interface of 15–20 and 20–25% Nycodenz. Four additional faint, light gray bands (bands 0, 1, 4, and 5) were visible at the interface of 0–10, 10–15, and 25–30% and in the pellet (Fig. 1A). Light microscopy analysis showed that bands 2 and 3 contained only etioplast-like organelles ranging in size between 2 and 4 μm (Fig. 1B). Differential interference contrast microscopy (Fig. 1C) revealed the etioplast-specific prolamellar body, suggesting that bands 2 and 3 contained highly purified etioplasts. Bands 0, 1, 4, and 5 (Fig. 1A) contained either etioplasts contaminated with other organelles or a mixture of different cell organelles and particles. Because light microscopy is not sufficient to exclude co-purification of other cell organelles during the Nycodenz density centrifugation, we analyzed the purity of etioplasts by comprehensive protein profiling using LC-MS/MS. We first identified diagnostic marker proteins from different cell organelles and the cytosol to monitor their presence in the different fractions obtained during the etioplast isolation procedure. Therefore we analyzed the supernatant of the first differential centrifugation and bands 0, 1, 4, and 5 (see Fig. 1A) from the first Nycodenz gradient by LC-MS/MS. Subsequently we analyzed the purified etioplasts isolated from band 2 of the second Nycodenz gradient and examined the protein complement of this band for the presence of marker proteins from other cell organelles. The purified etioplasts of bands 2 and 3 were later compared by differential display of their protein profiles using 2-D PAGE. For a valid comparison of the proteins and their relative abundance in the different gradient fractions and the supernatant, we applied the same protein shotgun identification strategy to all fractions. Although determination of protein abundance in complex LC-MS/MS experiments is only semiquantitative, conclusions about relative protein abundance can be drawn from a pairwise comparison of the number of different peptides identified from the same protein (i.e. sequence coverage) in two different samples (for a review, see Ref. 20Greenbaum D. Colangelo C. Williams K. Gerstein M. Comparing protein abundance and mRNA expression levels on a genomic scale.Genome Biol. 2003; 4: 117Google Scholar). Furthermore only those diagnostic marker proteins that are highly abundant in other cell compartments would be detected as contaminations in the etioplast fraction. Proteins that are enriched in the etioplast fraction compared with the other gradient fractions therefore can be considered true etioplast proteins or proteins that are specifically associated with the etioplast. This type of semiquantitative analysis is similar to a recently published protein profiling procedure using isotope tagging of membrane proteins (LOPIT (21Dunkley T.P. Watson R. Griffin J.L. Dupree P. Lilley K.S. Localization of organelle proteins by isotope tagging (LOPIT).Mol. Cell. Proteomics. 2004; 3: 1128-1134Google Scholar)). Both approaches allow a direct quantitative comparison of proteins from different cell organelles in the homologues system. This way, the enrichment of proteins by an organelle isolation procedure can be monitored. Together the described comparison of protein quantities in different protein fractions is more accurate to establish potential contaminations than cross-comparisons to different proteome studies that were performed with different biological material. We identified 725 proteins from all gradient fractions and the supernatant. Two hundred and forty different proteins were identified from etioplasts in band 2 (Supplemental Table 1, A and B), and 579 were identified from all other fractions (94 identified proteins of all other fractions were overlapping with band 2). All identified proteins were examined for their subcellular localization. We accepted those proteins as true plastid proteins that (i) are encoded in the plastid genome or have a predicted plastid transit peptide (22Emanuelsson O. Nielsen H. Brunak S. von Heijne G. Predicting subcellular localization of proteins based on their N-terminal amino acid sequence.J. Mol. Biol. 2000; 300: 1005-1016Google Scholar), (ii) are annotated as plastid proteins in the TIGR data base, or (iii) are homologues of chloroplast proteins as annotated in the