Chloroplast proteomics highlights the subcellular compartmentation of lipid metabolism

Progress in Lipid Research

Volumn 49, Issue 2, 2010, Pages 128-158

  Joyard, Jacques ^a   Ferro, Myriam ^b   Masselon, Christophe ^b   Seigneurin Berny, Daphné ^a   Salvi, Daniel ^a   Garin, Jérôme ^b   Rolland, Norbert ^a  

^a CEA GRENOBLE   (France)

^b INSERM   (France)

Author keywords

Envelope; Fatty acid desaturation; Fatty acid synthesis; Galactolipids; Glycerolipid synthesis; Stroma; Thylakoids

Indexed keywords

ACETYL COENZYME A; ACETYL COENZYME A CARBOXYLASE; DIACYLGLYCEROL; FRUCTOSE 1,6 BISPHOSPHATE; FRUCTOSE BISPHOSPHATE ALDOLASE; GALACTOLIPID; GLYCERALDEHYDE 3 PHOSPHATE DEHYDROGENASE; GLYCERALDEHYDE 3 PHOSPHATE DEHYDROGENASE A1; GLYCERALDEHYDE 3 PHOSPHATE DEHYDROGENASE A2; GLYCERALDEHYDE 3 PHOSPHATE DEHYDROGENASE B; GLYCEROLIPID; GLYCEROPHOSPHATE; PHOSPHATIDATE PHOSPHATASE; PHOSPHATIDIC ACID; PHOSPHOGLYCERATE KINASE; PHOSPHOGLYCERATE KINASE 1; PHOSPHOGLYCERATE KINASE 2; PHOSPHORIBULOKINASE; PPHOSPHORIBULOKINASE 1; PPHOSPHORIBULOKINASE 2; PYRUVATE DEHYDROGENASE; PYRUVIC ACID; RIBULOSEBISPHOSPHATE CARBOXYLASE; RIBULOSEBISPHOSPHATE CARBOXYLASE SMALL CHAIN 3B; TRANSALDOLASE; TRANSKETOLASE; TRANSKETOLASE 1; TRANSKETOLASE 2; UNCLASSIFIED DRUG; URIDINE DIPHOSPHATE GALACTOSE;

ACCURACY; ARABIDOPSIS; CELL COMPARTMENTALIZATION; CHLOROPLAST; FATTY ACID DESATURATION; FATTY ACID SYNTHESIS; FATTY ACID TRANSPORT; LIPID METABOLISM; NONHUMAN; PHENOTYPE; PHOTOSYNTHESIS; PLANT LEAF; PLANT SEED; PRIORITY JOURNAL; PROTEIN ANALYSIS; PROTEIN EXPRESSION; PROTEIN FUNCTION; PROTEIN LOCALIZATION; PROTEIN STRUCTURE; PROTEOMICS; REVIEW;

ARABIDOPSIS PROTEINS; BIOSYNTHETIC PATHWAYS; CELL MEMBRANE; CHLOROPLASTS; DATABASES, PROTEIN; FATTY ACIDS; LIPID METABOLISM; LIPIDS; PROTEOMICS; SUBCELLULAR FRACTIONS;

ARABIDOPSIS THALIANA; EUKARYOTA; PROKARYOTA;

EID: 76749131594     PISSN: 01637827     EISSN: None     Source Type: Journal    
DOI: 10.1016/j.plipres.2009.10.003     Document Type: Review

References (239)

1. Douce R., and Joyard J. Plant galactolipids. In: Stumpf P.K., and Conn E.E. (Eds). The biochemistry of plants. Lipids: structure and function vol. 4 (1980), Academic Press, New York 321-362
2. Siebertz H.P., Heinz E., Joyard J., and Douce R. Labelling in vivo and in vitro of molecular species of lipids from chloroplast envelopes and thylakoids. Eur J Biochem 108 (1980) 177-185
3. Heinz E. Enzymatic reactions in galactolipid biosynthesis. In: Tevini M., and Lichtenthaler H.K. (Eds). Lipids and lipid polymers (1977), Springer-Verlag, Berlin 102-120
4. Haverkate F., and van Deenen L.L. Isolation and chemical characterization of phosphatidyl glycerol from spinach leaves. Biochim Biophys Acta 106 (1965) 78-92
5. Dorne A.J., Joyard J., Block M.A., and Douce R. Localization of phosphatidylcholine in outer envelope membrane of spinach chloroplasts. J Cell Biol 100 (1985) 1690-1697
6. Dorne A.J., Joyard J., and Douce R. Do thylakoids really contain phosphatidylcholine?. Proc Natl Acad Sci USA 87 (1990) 71-74
7. Frentzen M. Phosphatidylglycerol and sulfoquinovosyldiacylglycerol: anionic membrane lipids and phosphate regulation. Curr Opin Plant Biol 7 (2004) 270-276
8. Jones M.R. Lipids in photosynthetic reaction centres: structural roles and functional holes. Prog Lipid Res 46 (2007) 56-87
9. Hölzl G., Witt S., Gaude N., Melzer M., Schöttler M.A., and Dörmann P. The role of diglycosyl lipids in photosynthesis and membrane lipid homeostasis in Arabidopsis. Plant Physiol 150 (2009) 1147-1159
10. Nussberger S., Dörr K., Wang D.N., and Kühlbrandt W. Lipid-protein interactions in crystals of plant light-harvesting complex. J Mol Biol 234 (1993) 347-356
11. Yu B., and Benning C. Anionic lipids are required for chloroplast structure and function in Arabidopsis. Plant J 36 (2003) 762-770
12. Pineau B., Girard-Bascou J., Eberhard S., Choquet Y., Trémolières A., Gérard-Hirne C., et al. A single mutation that causes phosphatidylglycerol deficiency impairs synthesis of photosystem II cores in Chlamydomonas reinhardtii. Eur J Biochem 271 (2004) 329-338
13. Dörmann P., Hoffmann-Benning S., Balbo I., and Benning C. Isolation and characterization of an Arabidopsis mutant deficient in the thylakoid lipid digalactosyl diacylglycerol. Plant Cell 7 (1995) 1801-1810
14. Härtel H., Lokstein H., Dörmann P., Grimm B., and Benning C. Changes in the composition of the photosynthetic apparatus in the galactolipid-deficient dgd1 mutant of Arabidopsis thaliana. Plant Physiol 115 (1997) 1175-1184
15. Jouhet J., Maréchal E., Baldan B., Bligny R., Joyard J., and Block M.A. Phosphate deprivation induces transfer of DGDG galactolipid from chloroplast to mitochondria. J Cell Biol 167 (2004) 863-874
16. Blée E. Impact of phyto-oxylipins in plant defense. Trends Plant Sci 7 (2002) 315-322
17. Andreou A., Brodhun F., and Feussner I. Biosynthesis of oxylipins in non-mammals. Prog Lipid Res 48 (2009) 148-170
18. Slabas A.R., and Fawcett T. The biochemistry and molecular biology of plant lipid biosynthesis. Plant Mol Biol 19 (1992) 169-191
19. Harwood J.L. Recent advances in the biosynthesis of plant fatty acids. Biochim Biophys Acta 1301 (1996) 7-56
20. Joyard J., and Douce R. Galactolipid synthesis. In: Stumpf P.K. (Ed). The biochemistry of plants. Lipids: structure and function vol. 9 (1987), Academic Press, New York 215-274
21. Bates P.D., Ohlrogge J.B., and Pollard M. Incorporation of newly synthesized fatty acids into cytosolic glycerolipids in pea leaves occurs via acyl editing. J Biol Chem 282 (2007) 31206-31216
22. Browse J., Warwick N., Somerville C.R., and Slack C.R. Fluxes through the prokaryotic and eukaryotic pathways of lipid synthesis in the 16:3 plant Arabidopsis thaliana. Biochem J 235 (1986) 25-31
23. Jouhet J., Maréchal E., and Block M.A. Glycerolipid transfer for the building of membranes in plant cells. Prog Lipid Res 46 (2007) 37-55
24. Benning C., Xu C., and Awai K. Non-vesicular and vesicular lipid trafficking involving plastids. Curr Opin Plant Biol 9 (2006) 241-247
25. Benning C. A role for lipid trafficking in chloroplast biogenesis. Prog Lipid Res 47 (2008) 381-389
26. Benning C. Mechanism of lipid transport involved in organelle biogenesis in plant cells. Annu Rev Cell Dev Biol 25 (2009) 71-91
27. Stumpf P.K. Metabolism of fatty acids. Annu Rev Biochem 38 (1969) 159-212
28. Joyard J., and Douce R. Site of synthesis of phosphatidic acid and diacylglycerol in spinach chloroplasts. Biochim Biophys Acta 486 (1977) 273-285
29. Roughan P.G., and Slack C.R. Cellular organization of glycerolipid metabolism. Annu Rev Plant Physiol 33 (1982) 97-132
30. Heinz E., and Roughan P.G. Similarities and differences in lipid metabolism of chloroplasts isolated from 18:3 and 16:3 plants. Plant Physiol 72 (1983) 273-279
31. Slabas A.R., Brown A., Sinden B.S., Swinhoe R., Simon J.W., Ashton A.R., et al. Pivotal reactions in fatty acid synthesis. Prog Lipid Res 33 (1994) 39-46
32. Heinz E. Plant glycolipids. In: Christie W.W. (Ed). Advances in lipid methodology - three (1996), The Oily Press, Dundee 211-332 [chapter 6]
33. Ohlrogge J., and Browse J. Lipid biosynthesis. Plant Cell 7 (1995) 957-970
34. Ohlrogge J.B., and Jaworski J.G. Regulation of fatty acid synthesis. Annu Rev Plant Physiol Plant Mol Biol 48 (1997) 109-136
35. Awai K., Maréchal E., Block M.A., Brun D., Masuda T., Shimada H., et al. Two types of MGDG synthase genes, found widely in both 16:3 and 18:3 plants, differentially mediate galactolipid syntheses in photosynthetic and nonphotosynthetic tissues in Arabidopsis thaliana. Proc Natl Acad Sci USA 98 (2001) 10960-10965
36. Somerville C., and Browse J. Plant lipids: metabolism, mutants, and membranes. Science 252 (1991) 80-87
37. Arondel V., Lemieux B., Hwang I., Gibson S., Goodman H.M., and Somerville C.R. Map-based cloning of a gene controlling omega-3 fatty acid desaturation in Arabidopsis. Science 258 (1992) 1353-1355
38. Shimojima M., Ohta H., Iwamatsu A., Masuda T., Shioi Y., and Takamiya K. Cloning of the gene for monogalactosyldiacylglycerol synthase and its evolutionary origin. Proc Natl Acad Sci USA 94 (1997) 333-337
39. Mekhedov S., de Ilárduya O.M., and Ohlrogge J. Toward a functional catalog of the plant genome. A survey of genes for lipid biosynthesis. Plant Physiol 122 (2000) 389-402
40. Dörmann P., and Benning C. Galactolipids rule in seed plants. Trends Plant Sci 7 (2002) 112-118
41. Wallis J.G., and Browse J. Mutants of Arabidopsis reveal many roles for membrane lipids. Prog Lipid Res 41 (2002) 254-278
42. Beisson F., Koo A.J., Ruuska S., Schwender J., Pollard M., Thelen J.J., et al. Arabidopsis thaliana genes involved in acyl lipid metabolism. A 2003 census of the candidates, a study of the distribution of expressed sequence tags in organs, and a web-based database. Plant Physiol 132 (2003) 681-697
43. Sun Q., Zybailov B., Majeran W., Friso G., Olinares P.D., and van Wijk K.J. PPDB, the plant proteomics database at cornell. Nucleic Acid Res 37 (2009) D969-D974
44. Heazlewood J.L., Verboom R.E., Tonti-Filippini J., Small I., and Millar A.H. SUBA: the Arabidopsis Subcellular Database. Nucleic Acid Res 35 D (2007) 213-218
45. Ferro M, Brugière S, Salvi D, Seigneurin-Berny D, Court C, Moyet L, et al. AT_CHLORO: a comprehensive chloroplast proteome database with sub-plastidial localization and information for functional genomics using quantitative label-free analyses (submitted for publication).
46. Ferro M., Salvi D., Rivière-Rolland H., Vermat T., Seigneurin-Berny D., Grunwald D., et al. Integral membrane proteins of the chloroplast envelope: identification and subcellular localization of new transporters. Proc Natl Acad Sci USA 99 (2002) 11487-11492
47. Ferro M., Salvi D., Brugière S., Miras S., Kowalski S., Louwagie M., et al. Proteomics of the chloroplast envelope membranes from Arabidopsis thaliana. Mol Cell Proteomics 2 (2003) 325-345
48. Friso G., Ytterberg A.J., Giacomelli L., Peltier J.B., Rudella A., Sun Q., et al. In-depth analysis of the thylakoid membrane proteome of Arabidopsis thaliana chloroplasts; new proteins, functions and a plastid proteome database. Plant Cell 16 (2004) 478-499
49. Froehlich J.E., Wilkerson C.G., Ray W.K., McAndrew R.S., Osteryoung K.W., Gage D.A., et al. Proteomic study of the Arabidopsis thaliana chloroplastic envelope membrane utilizing alternatives to traditional two-dimensional electrophoresis. J Proteome Res 2 (2003) 413-425
50. Kleffmann T., Russenberger D., von Zychlinski A., Christopher W., Sjölander K., Gruissem W., et al. The Arabidopsis thaliana chloroplast proteome reveals pathway abundance and novel protein functions. Curr Biol 14 (2004) 354-362
51. Peltier J.B., Emanuelsson O., Kalume D.E., Ytterberg J., Friso G., Rudella A., et al. Central functions of the lumenal and peripheral thylakoid proteome of Arabidopsis determined by experimentation and genome-wide prediction. Plant Cell 14 (2002) 211-236
52. Peltier J.B., Ytterberg A.J., Sun Q., and 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 279 (2004) 49367-49383
53. Peltier J.B., Cai Y., Sun Q., Zabrouskov V., Giacomelli L., Rudella A., et al. The oligomeric stromal proteome of Arabidopsis thaliana chloroplasts. Mol Cell Proteomics 5 (2006) 114-133
54. Majeran W., Zybailov B., Ytterberg A.J., Dunsmore J., Sun Q., and van Wijk K.J. Consequences of C4 differentiation for chloroplast membrane proteomes in maize mesophyll and bundle sheath cells. Mol Cell Proteomics 7 (2008) 1609-1638
55. Zybailov B., Rutschow H., Friso G., Rudella A., Emanuelsson O., Sun Q., et al. Sorting signals, N-terminal modifications and abundance of the chloroplast proteome. PLoS ONE 3 (2008) e1994
56. Rhee S.Y., Beavis W., Berardini T.Z., Chen G., Dixon D., Doyle A., et al. The Arabidopsis Information Resource (TAIR): a model organism database providing a centralized, curated gateway to Arabidopsis biology, research materials and community. Nucleic Acid Res 31 (2003) 224
57. Emanuelsson O., Nielsen H., von Heijne G., and ChloroP. a neural network-based method for predicting chloroplast transit peptides and their cleavage sites. Protein Sci 8 (1999) 978-984
58. Emanuelsson O., Brunak S., von Heijne G., and Nielsen H. Locating proteins in the cell using TargetP, SignalP, and related tools. Nat Protocols 2 (2007) 953-971
59. Schwacke R., Schneider A., Van Der Graaff E., Fischer K., Catoni E., Desimone M., et al. ARAMEMNON, a novel database for Arabidopsis integral membrane proteins. Plant Physiol 131 (2003) 16-26
60. Heazlewood J.L., Tonti-Filippini J.S., Gout A.M., Day D.A., Whelan J., and Millar A.H. Experimental analysis of the Arabidopsis mitochondrial proteome highlights signaling and regulatory components, provides assessment of targeting prediction programs, and indicates plant-specific mitochondrial proteins. Plant Cell 16 (2004) 241-256
61. Ströher E., and Dietz K.J. The dynamic thiol-disulphide redox proteome of the Arabidopsis thaliana chloroplast as revealed by differential electrophoretic mobility. Physiol Plant 133 (2008) 566-583
62. Goulas E., Schubert M., Kieselbach T., Kleczkowski L.A., Gardeström P., Schröder W., et al. The chloroplast lumen and stromal proteomes of Arabidopsis thaliana show differential sensitivity to short- and long-term exposure to low temperature. Plant J 47 (2006) 720-734
63. Bae M.S., Cho E.J., Choi E.Y., and Park O.K. Analysis of the Arabidopsis nuclear proteome and its response to cold stress. Plant J 36 (2003) 652-663
64. Giacomelli L., Rudella A., and van Wijk K.J. High light response of the thylakoid proteome in Arabidopsis wild type and the ascorbate-deficient mutant vtc2-2. A comparative proteomics study. Plant Physiol 141 (2006) 685-701
65. Ytterberg A.J., Peltier J.B., and van Wijk K.J. Protein profiling of plastoglobules in chloroplasts and chromoplasts. A surprising site for differential accumulation of metabolic enzymes. Plant Physiol 140 (2006) 984-997
66. Vidi P.A., Kanwischer M., Baginsky S., Austin J.R., Csucs G., Dörmann P., et al. Tocopherol cyclase (VTE1) localization and vitamin E accumulation in chloroplast plastoglobule lipoprotein particles. J Biol Chem 281 (2006) 11225-11234
67. Schubert M., Petersson U.A., Haas B.J., Funk C., Schröder W.P., and Kieselbach T. Proteome map of the chloroplast lumen of Arabidopsis thaliana. J Biol Chem 277 (2002) 8354-8365
68. Lee C.P., Eubel H., O'Toole N., and Millar A.H. Heterogeneity of the mitochondrial proteome for photosynthetic and non-photosynthetic Arabidopsis metabolism. Mol Cell Proteomics 7 (2008) 1297-1316
69. Pendle A.F., Clark G.P., Boon R., Lewandowska D., Lam Y.W., Andersen J., et al. Proteomic analysis of the Arabidopsis nucleolus suggests novel nucleolar functions. Mol Biol Cell 16 (2005) 260-269
70. Marmagne A., Ferro M., Meinnel T., Bruley C., Kuhn L., Garin J., et al. A high content in lipid-modified peripheral proteins and integral receptor kinases features in the Arabidopsis plasma membrane proteome. Mol Cell Proteomics 6 (2007) 1980-1996
71. Sweetlove L.J., Heazlewood J.L., Herald V., Holtzapffel R., Day D.A., Leaver C.J., et al. The impact of oxidative stress on Arabidopsis mitochondria. Plant J 32 (2002) 891-904
72. Benschop J.J., Mohammed S., O'Flaherty M., Heck A.J., Slijper M., and Menke F.L. Quantitative phosphoproteomics of early elicitor signaling in Arabidopsis. Mol Cell Proteomics 6 (2007) 1198-1214
73. Jaquinod M., Villiers F., Kieffer-Jaquinod S., Hugouvieux V., Bruley C., Garin J., et al. A proteomics dissection of Arabidopsis thaliana vacuoles isolated from cell culture. Mol Cell Proteomics 6 (2007) 394-412
74. Alexandersson E., Saalbach G., Larsson C., and Kjellbom P. Arabidopsis plasma membrane proteomics identifies components of transport, signal transduction and membrane trafficking. Plant Cell Physiol 45 (2004) 1543-1556
75. Boudart G., Jamet E., Rossignol M., Lafitte C., Borderies G., Jauneau A., et al. Cell wall proteins in apoplastic fluids of Arabidopsis thaliana rosettes: identification by mass spectrometry and bioinformatics. Proteomics 5 (2005) 212-221
76. Dunkley T.P., Hester S., Shadforth I.P., Runions J., Weimar T., Hanton S.L., et al. Mapping the Arabidopsis organelle proteome. Proc Natl Acad Sci USA 103 (2006) 6518-6523
77. Fukao Y., Hayashi M., Hara-Nishimura I., and Nishimura M. Novel glyoxysomal protein kinase, GPK1, identified by proteomic analysis of glyoxysomes in etiolated cotyledons of Arabidopsis thaliana. Plant Cell Physiol 44 (2003) 1002-1012
78. Ito J., Heazlewood J.L., and Millar A.H. Analysis of the soluble ATP-binding proteome of plant mitochondria identifies new proteins and nucleotide triphosphate interactions within the matrix. J Proteome Res 5 (2006) 3459-3469
79. Millar A.H., Sweetlove L.J., Giegé P., and Leaver C.J. Analysis of the Arabidopsis mitochondrial proteome. Plant Physiol 127 (2001) 1711-1727
80. Kruft V., Eubel H., Jänsch L., Werhahn W., and Braun H.P. Proteomic approach to identify novel mitochondrial proteins in Arabidopsis. Plant Physiol 127 (2001) 1694-1710
81. Reumann S., Quan S., Aung K., Yang P., Manandhar-Shrestha K., Holbrook D., et al. In-depth proteome analysis of Arabidopsis leaf peroxisomes combined with in vivo subcellular targeting verification indicates novel metabolic and regulatory functions of peroxisomes. Plant Physiol 150 (2009) 125-143
82. Reumann S., Babujee L., Ma C., Wienkoop S., Siemsen T., Antonicelli G.E., et al. Proteome analysis of Arabidopsis leaf peroxisomes reveals novel targeting peptides, metabolic pathways, and defense mechanisms. Plant Cell 19 (2007) 3170-3193
83. Eubel H., Meyer E.H., Taylor N.L., Bussell J.D., O'Toole N., Heazlewood J.L., et al. Novel proteins, putative membrane transporters, and an integrated metabolic network are revealed by quantitative proteomic analysis of Arabidopsis cell culture peroxisomes. Plant Physiol 148 (2008) 1809-1829
84. Slabas A.R., Ndimba B., Simon W.J., and Chivasa S. Proteomic analysis of the Arabidopsis cell wall reveals unexpected proteins with new cellular locations. Biochem Soc Trans 32 (2004) 524-528
85. Szponarski W., Sommerer N., Boyer J.C., Rossignol M., and Gibrat R. Large-scale characterization of integral proteins from Arabidopsis vacuolar membrane by two-dimensional liquid chromatography. Proteomics 4 (2004) 397-406
86. Jamet E., Canut H., Boudart G., and Pont-Lezica R.F. Cell wall proteins: a new insight through proteomics. Trends Plant Sci 11 (2006) 33-39
87. Brugière S., Kowalski S., Ferro M., Seigneurin-Berny D., Miras S., Salvi D., et al. The hydrophobic proteome of mitochondrial membranes from Arabidopsis cell suspensions. Phytochemistry 65 (2004) 1693-1707
88. Nelson C.J., Hegeman A.D., Harms A.C., and Sussman M.R. A quantitative analysis of Arabidopsis plasma membrane using trypsin-catalyzed (18)O labeling. Mol Cell Proteomics 5 (2006) 1382-1395
89. Nühse T.S., Stensballe A., Jensen O.N., and Peck S.C. Phosphoproteomics of the Arabidopsis plasma membrane and a new phosphorylation site database. Plant Cell 16 (2004) 2394-2405
90. Givan C.V. The source of acetyl coenzyme A in chloroplasts of higher plants. Physiol Plant 57 (1983) 311-316
91. Bao X., Focke M., Pollard M., and Ohlrogge J. Understanding in vivo carbon precursor supply for fatty acid synthesis in leaf tissue. Plant J 22 (2000) 39-50
92. Smirnov B.P. The biosynthesis of higher fatty acids from acetate in isolated chloroplasts from Spinacia oleracea. Biokhimiya 25 (1960) 545-555
93. Roughan P.G., and Ohlrogge J.B. Evidence that isolated chloroplasts contain an integrated lipid-synthesizing assembly that channels acetate into long-chain fatty acids. Plant Physiol 110 (1996) 1239-1247
94. Ruuska S.A., Schwender J., and Ohlrogge J.B. The capacity of green oilseeds to utilize photosynthesis to drive biosynthetic processes. Plant Physiol 136 (2004) 2700-2709
95. Schwender J., Goffman F., Ohlrogge J.B., and Shachar-Hill Y. Rubisco without the Calvin cycle improves the carbon efficiency of developing green seeds. Nature 432 (2004) 779-782
96. Ke J., Wen T.N., Wurtele E.S., and Nikolau B.J. Coordinate regulation of the spatial and temporal expression of the nuclear and chloroplastic encoded genes of the heteromeric acetyl-CoA carboxylase. Plant Physiol 122 (2000) 1057-1071
97. Andre C., Froehlich J.E., Moll M.R., and Benning C. A heteromeric plastidic pyruvate kinase complex involved in seed oil biosynthesis in Arabidopsis. Plant Cell 19 (2007) 2006-2022
98. Mentzen W.I., Peng J., Ransom N., Nikolau B.J., and Wurtele E.S. Articulation of three core metabolic processes in Arabidopsis: fatty acid biosynthesis, leucine catabolism and starch metabolism. BMC Plant Biol 8 (2008) 76
99. Irving L.J., and Robinson D. A dynamic model of Rubisco turnover in cereal leaves. New Phytol 169 (2006) 493-504
100. Fischer K., and Weber A. Transport of carbon in non-green plastids. Trends Plant Sci 7 (2002) 345-351
101. Li H., Culligan K., Dixon R.A., and Chory J. CUE1: a mesophyll cell-specific positive regulator of light-controlled gene expression in Arabidopsis. Plant Cell 7 (1995) 1599-1610
102. Streatfield S.J., Weber A., Kinsman E.A., Häusler R.E., Li J., Post-Beittenmiller D., et al. The phosphoenolpyruvate/phosphate translocator is required for phenolic metabolism, palisade cell development, and plastid-dependent nuclear gene expression. Plant Cell 11 (1999) 1609-1622
103. Shen J., Ren X., Cao R., Liu J., and Gong Z. Transcriptional gene silencing mediated by a plastid inner envelope Phosphoenolpyruvate/Phosphate Translocator CUE1 in Arabidopsis. Plant Physiol 150 (2009) 1990-1996
104. Lin M., and Oliver D.J. The role of acetyl-coenzyme A synthetase in Arabidopsis. Plant Physiol 147 (2008) 1822-1829
105. Leustek T., and Saito K. Sulfate transport and assimilation in plants. Plant Physiol 120 (1999) 637-643
106. Kürsteiner O., Dupuis I., and Kuhlemeier C. The pyruvate decarboxylase1 gene of Arabidopsis is required during anoxia but not other environmental stresses. Plant Physiol 132 (2003) 968-978
107. Mooney B.P., Miernyk J.A., and Randall D.D. The complex fate of alpha-ketoacids. Annu Rev Plant Biol 53 (2002) 357-375
108. Tovar-Méndez A., Miernyk J.A., and Randall D.D. Regulation of pyruvate dehydrogenase complex activity in plant cells. Eur J Biochem 270 (2003) 1043-1049
109. Cronan Jr. J.E., and Waldrop G.L. Multi-subunit acetyl-CoA carboxylases. Prog Lipid Res 41 (2002) 407-435
110. Sasaki Y., and Nagano Y. Plant acetyl-CoA carboxylase: structure, biosynthesis, regulation, and gene manipulation for plant breeding. Biosci Biotechnol Biochem 68 (2004) 1175-1184
111. Thelen J.J., and Ohlrogge J.B. The multisubunit acetyl-CoA carboxylase is strongly associated with the chloroplast envelope through non-ionic interactions to the carboxyltransferase subunits. Arch Biochem Biophys 400 (2002) 245-257
112. Shintani D., Roesler K., Shorrosh B., Savage L., and Ohlrogge J. Antisense expression and overexpression of biotin carboxylase in tobacco leaves. Plant Physiol 114 (1997) 881-886
113. Chen M., Mooney B.P., Hajduch M., Joshi T., Zhou M., Xu D., et al. System analysis of an Arabidopsis mutant altered in de novo fatty acid synthesis reveals diverse changes in seed composition and metabolism. Plant Physiol 150 (2009) 27-41
114. Guerra D.J., Ohlrogge J.B., and Frentzen M. Activity of acyl carrier protein isoforms in reactions of plant fatty acid metabolism. Plant Physiol 82 (1986) 448-453
115. Pidkowich M.S., Nguyen H.T., Heilmann I., Ischebeck T., and Shanklin J. Modulating seed beta-ketoacyl-acyl carrier protein synthase II level converts the composition of a temperate seed oil to that of a palm-like tropical oil. Proc Natl Acad Sci USA 104 (2007) 4742-4747
116. Mou Z., He Y., Dai Y., Liu X., and Li J. Deficiency in fatty acid synthase leads to premature cell death and dramatic alterations in plant morphology. Plant Cell 12 (2000) 405-418
117. Serrano M., Robatzek S., Torres M., Kombrink E., Somssich I.E., Robinson M., et al. Chemical interference of pathogen-associated molecular pattern-triggered immune responses in Arabidopsis reveals a potential role for fatty-acid synthase type II complex-derived lipid signals. J Biol Chem 282 (2007) 6803-6811
118. Dayan F.E., Ferreira D., Wang Y.H., Khan I.A., McInroy J.A., and Pan Z. A pathogenic fungi diphenyl ether phytotoxin targets plant enoyl (acyl carrier protein) reductase. Plant Physiol 147 (2008) 1062-1071
119. Mu J., Tan H., Zheng Q., Fu F., Liang Y., Zhang J., et al. Leafy cotyledon1 is a key regulator of fatty acid biosynthesis in Arabidopsis. Plant Physiol 148 (2008) 1042-1054
120. Shanklin J., and Cahoon E.B. Desaturation and related modification of fatty acids. Annu Rev Plant Physiol Plant Mol Biol 49 (1998) 611-641
121. Lindqvist Y., Huang W., Schneider G., and Shanklin J. Crystal structure of delta9 stearoyl-acyl carrier protein desaturase from castor seed and its relationship to other di-iron proteins. EMBO J 15 (1996) 4081-4092
122. Kachroo A., Shanklin J., Whittle E., Lapchyk L., Hildebrand D., and Kachroo P. The Arabidopsis stearoyl-acyl carrier protein-desaturase family and the contribution of leaf isoforms to oleic acid synthesis. Plant Mol Biol 63 (2007) 257-271
123. Kachroo A., Lapchyk L., Fukushige H., Hildebrand D., Klessig D., and Kachroo P. Plastidial fatty acid signaling modulates salicylic acid- and jasmonic acid-mediated defense pathways in the Arabidopsis ssi2 mutant. Plant Cell 15 (2003) 2952-2965
124. Rawsthorne S. Carbon flux and fatty acid synthesis in plants. Prog Lipid Res 41 (2002) 182-196
125. Voelker T., and Kinney A.J. Variations in the biosynthesis of seed storage lipids. Annu Rev Plant Physiol Plant Mol Biol 52 (2001) 335-361
126. Salas J.J., and Ohlrogge J.B. Characterization of substrate specificity of plant FatA and FatB acyl-ACP thioesterases. Arch Biochem Biophys 403 (2002) 25-34
127. Mayer K.M., and Shanklin J. A structural model of the plant acyl-acyl carrier protein thioesterase FatB comprises two helix/4-stranded sheet domains, the N-terminal domain containing residues that affect specificity and the C-terminal domain containing catalytic residues. J Biol Chem 280 (2005) 3621-3627
128. Joyard J., and Stumpf P.K. Synthesis of long-chain Acyl-CoA in chloroplast envelope membranes. Plant Physiol 67 (1981) 250-256
129. Block M.A., Dorne A.J., Joyard J., and Douce R. The acyl-CoA synthetase and the acyl-CoA thioesterase are located respectively on the outer and on the inner membrane of the chloroplast envelope. FEBS Lett 153 (1983) 377-381
130. Andrews J., and Keegstra K. Acyl-CoA synthetase is located in the outer membrane and Acyl-CoA thioesterase in the inner membrane of Pea chloroplast envelopes. Plant Physiol 72 (1983) 735-740
131. Koo A.J., Ohlrogge J.B., and Pollard M. On the export of fatty acids from the chloroplast. J Biol Chem 279 (2004) 16101-16110
132. Shockey J.M., Fulda M.S., and Browse J.A. Arabidopsis contains nine long-chain acyl-coenzyme A synthetase genes that participate in fatty acid and glycerolipid metabolism. Plant Physiol 129 (2002) 1710-1722
133. Schnurr J.A., Shochey J.M., de Boer G.J., and Browse J.A. Fatty acid export from the chloroplast. Molecular characterization of a major plastidial acyl-coenzyme A synthetase from Arabidopsis. Plant Physiol 129 (2002) 1700-1709
134. Koo A.J., Fulda M., Browse J., and Ohlrogge J.B. Identification of a plastid acyl-acyl carrier protein synthetase in Arabidopsis and its role in the activation and elongation of exogenous fatty acids. Plant J 44 (2005) 620-632
135. Rea P.A. Plant ATP-binding cassette transporters. Annu Rev Plant Biol 58 (2007) 347-375
136. Koo A.J., and Ohlrogge J.B. The predicted candidates of Arabidopsis plastid inner envelope membrane proteins and their expression profiles. Plant Physiol 130 (2002) 823-836
137. Weber A.P., Schwacke R., and Flügge U.I. Solute transporters of the plastid envelope membrane. Annu Rev Plant Biol 56 (2005) 133-164
138. Miquel M., Cassagne C., and Browse J. A new class of Arabidopsis mutants with reduced hexadecatrienoic acid fatty acid levels. Plant Physiol 117 (1998) 923-930
139. Nandi A., Welti R., and Shah J. The Arabidopsis thaliana dihydroxyacetone phosphate reductase gene suppresssor of fatty acid desaturase deficiency1 is required for glycerolipid metabolism and for the activation of systemic acquired resistance. Plant Cell 16 (2004) 465-477
140. Frentzen M., Heinz E., McKeon T.A., and Stumpf P.K. Specificities and selectivities of glycerol-3-phosphate acyltransferase and monoacylglycerol-3-phosphate acyltransferase from pea and spinach chloroplasts. Eur J Biochem 129 (1983) 629-636
141. Zheng Z., Xia Q., Daux M., Shen W., Selvaraj G., and Zou J. Arabidopsis AtGPAT 1, a member of the membrane-bound glycerol-3-phosphate acyltransferase gene family, is essential to tapetum differentiation and male fertility. Plant Cell 15 (2003) 1872-1887
142. Gidda S.K., Shockey J.M., Rothstein S.J., Dyer J.M., and Mullen R.T. Arabidopsis thaliana GPAT 8 and GPAT 9 are localized to the ER and possess distinct ER retrieval signals: functional divergence of the dilysine ER retrieval motif in plant cells. Plant Physiol Biochem 47 (2009) 867-879
143. Kim H.U., and Huang A.H.C. Plastid lysophosphatidyl acyltransferase is essential for embryo development in Arabidopsis. Plant Physiol 134 (2004) 1206-1216
144. Turnbull A.P., Rafferty J.B., Sedelnikova S.E., Slabas A.R., Schierer T.P., Kroon J.T., et al. Analysis of the structure, substrate specificity, and mechanism of squash glycerol-3-phosphate (1)-acyltransferase. Structure 9 (2001) 347-353
145. Tamada T., Feese M.D., Ferri S.R., Kato Y., Yajima R., Toguri T., et al. Substrate recognition and selectivity of plant glycerol-3-phosphate acyltransferases (GPATs) from Cucurbita moscata and Spinacea oleracea. Acta Crystallogr D60 (2004) 13-21
146. Kunst L., Browse J., and Somerville C.R. Altered regulation of lipid biosynthesis in a mutant of Arabidopsis deficient in chloroplast glycerol-3-phosphate acyltransferase activity. Proc Natl Acad Sci USA 85 (1988) 4143-4147
147. Fritz M., Lokstein H., Hackenberg D., Welti R., Roth M., Zähringer U., et al. Channeling of eukaryotic diacylglycerol into the biosynthesis of plastidial phosphatidylglycerol. J Biol Chem 282 (2007) 4613-4625
148. Xu C., Yu B., Cornish A.J., Froehlich J.E., and Benning C. Phosphatidylglycerol biosynthesis in chloroplasts of Arabidopsis mutants deficient in acyl-ACP glycerol-3- phosphate acyltransferase. Plant J 47 (2006) 296-309
149. Nishida I., Frentzen M., Ishizaki O., and Murata N. Purification of isomeric forms of acyl-(acyl-carrier-protein): glycerol-3-phosphate acyltransferase from greening squash cotyledons. Plant Cell Physiol 28 (1997) 1071-1079
150. Nishida I., Sugiura M., Enju A., and Nakamura M. A second gene for acyl-(acyl-carrier-protein): glycerol-3-phosphate acyltransferase in squash, Cucurbita moschata cv. Shirogikuza, codes for an oleate-selective isozyme: molecular cloning and protein purification studies. Plant Cell Physiol 41 (2000) 1381-1391
151. Yu B., Wakao S., Fan J., and Benning C. Loss of plastidic lysophosphatidic acid acyltransferase causes embryo-lethality in Arabidopsis. Plant Cell Physiol 45 (2004) 503-510
152. Joyard J., and Douce R. Characterisation of phosphatidate phosphohydrolase activity associated with the chloroplast envelope. FEBS Lett 102 (1979) 147-150
153. Malherbe A., Block M.A., Joyard J., and Douce R. Feedback inhibition of phosphatidate phosphatase from spinach chloroplast envelope membranes by diacylglycerol. J Biol Chem 267 (1992) 23546-23553
154. Block M.A., Dorne A.J., Joyard J., and Douce R. The phosphatidic acid phosphatase is located on the inner membrane of the spinach chloroplast envelope. FEBS Lett 164 (1983) 111-115
155. Andrews J., Ohlrogge J.B., and Keegstra K. Final step of phosphatidic acid synthesis in Pea chloroplasts occurs in the inner envelope membrane. Plant Physiol 78 (1985) 459-465
156. Nakamura Y., Tsuchiya M., and Ohta H. Plastidic phosphatidic acid phosphatases identified in a distinct subfamily of lipid phosphate phosphatases with prokaryotic origin. J Biol Chem 282 (2007) 29013-29021
157. Awai K., Xu C., Tamot B., and Benning C. A phosphatidic acid-binding protein of the chloroplast inner envelope membrane involved in lipid trafficking. Proc Natl Acad Sci USA 103 (2006) 10817-10822
158. Li M., Hong Y., and Wang X. Phospholipase d- and phosphatidic acid-mediated signaling in plants. Biochim Biophys Acta 1791 (2009) 927-935
159. Xu C., Fan J., Riekhof W., Froehlich J.E., and Benning C. A permease-like protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J 22 (2003) 2370-2379
160. Lu B., Xu C., Awai K., Jones A.D., and Benning C. A small ATPase protein of Arabidopsis, TGD3, involved in chloroplast lipid import. J Biol Chem 282 (2007) 35945-35953
161. Xu C., Fan J., Cornish A.J., and Benning C. Lipid trafficking between the endoplasmic reticulum and the plastid in Arabidopsis requires the extraplastidic TGD4 protein. Plant Cell 20 (2008) 2190-2204
162. Xu C., Fan J., Froehlich J.E., Awai K., and Benning C. Mutation of the TGD1 chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. Plant Cell 17 (2005) 3094-3110
163. Douce R., and Joyard J. Biochemistry and function of the plastid envelope. Annu Rev Cell Biol 6 (1990) 173-216
164. 31P NMR studies of spinach leaves and their chloroplasts. J Biol Chem 265 (1990) 1319-1326
165. Barber C., Rösti J., Rawat A., Findlay K., Roberts K., and Seifert G.J. Distinct properties of the five UDP-d-glucose/UDP-d-galactose 4-epimerase usoforms of Arabidopsis thaliana. J Biol Chem 281 (2006) 17276-17285
166. Dörmann P., and Benning C. The role of UDP-glucose epimerase in carbohydrate metabolism of Arabidopsis. Plant J 13 (1998) 641-652
167. Seifert G.J., Barber C., Wells B., Dolan L., and Roberts K. Galactose biosynthesis in Arabidopsis: genetic evidence for substrate channeling from UDP-d-galactose into cell wall polymers. Curr Biol 12 (2002) 1840-1845
168. Douce R. Site of biosynthesis of galactolipids in spinach chloroplasts. Science 183 (1974) 852-853
169. Carter H.E., McCluer R.H., and Slifer E.D. Lipids of wheat flour. I. Characterization of galactosylglycerol components. J Am Chem Soc 78
170. Miège C., Maréchal E., Shimojima M., Awai K., Block M.A., Ohta H., et al. Biochemical and topological properties of type A MGDG synthase, a spinach chloroplast envelope enzyme catalyzing the synthesis of both prokaryotic and eukaryotic MGDG. Eur J Biochem 265 (1999) 990-1001
171. Kobayashi K., Kondo M., Fukuda H., Nishimura M., and Ohta H. Galactolipid synthesis in chloroplast inner envelope is essential for proper thylakoid biogenesis, photosynthesis, and embryogenesis. Proc Natl Acad Sci USA 104 (2007) 17216-17221
172. Jarvis P., Dörmann P., Peto C.A., Lutes J., Benning C., and Chory J. Galactolipid deficiency and abnormal chloroplast development in the Arabidopsis MGD synthase 1 mutant. Proc Natl Acad Sci USA 97 (2000) 8175-8179
173. Kobayashi K., Awai K., Nakamura M., Nagatani A., Masuda T., and Ohta H. Type-B monogalactosyldiacylglycerol synthases are involved in phosphate starvation-induced lipid remodeling, and are crucial for low-phosphate adaptation. Plant J 57 (2009) 322-331
174. van Besouw A., and Wintermans J.F. Galactolipid formation in chloroplast envelopes. I. Evidence for two mechanisms in galactosylation. Biochim Biophys Acta 529 (1978) 44-53
175. Dorne A.J., Block M.A., Joyard J., and Douce R. The galactolipid:galactolipid galactosyltransferase is located on the outer surface of the outer chloroplast envelope. FEBS Lett 145 (1982) 30-34
176. Dörmann P., Balbo I., and Benning C. Arabidopsis galactolipid biosynthesis and lipid trafficking mediated by DGD1. Science 284 (1999) 2181-2184
177. Kelly A.A., and Dörmann P. DGD2, an Arabidopsis gene encoding a UDP-galactose-dependent digalactosyldiacylglycerol synthase is expressed during growth under phosphate-limiting conditions. J Biol Chem 277 (2002) 1166-1173
178. Froehlich J.E., Benning C., and Dörmann P. The digalactosyldiacylglycerol (DGDG) synthase DGD1 is inserted into the outer envelope membrane of chloroplasts in a manner independent of the general import pathway and does not depend on direct interaction with monogalactosyldiacylglycerol synthase for DGDG biosynthesis. J Biol Chem 276 (2001) 31806-31812
179. Kelly A.A., Froehlich J.E., and Dörmann P. Disruption of the two digalactosyldiacylglycerol synthase genes DGD1 and DGD2 in Arabidopsis reveals the existence of an additional enzyme of galactolipid synthesis. Plant Cell 15 (2003) 2694-2706
180. Misson J., Raghothama K.G., Jain A., Jouhet J., Block M.A., Bligny R., et al. A wide transcriptional analysis using the Arabidopsis thaliana whole genome Affymetrix gene chips determined plant responses to phosphate deprivation. Proc Natl Acad Sci USA 102 (2005) 11934-11939
181. Chen L.J., and Li H.M. A mutant deficient in the plastid lipid DGD is defective in protein import into chloroplasts. Plant J 16 (1998) 33-39
182. Sakaki T., Saito K., Kawaguchi A., Kondo N., and Yamada M. Conversion of monogalactosyldiacylglycerols to triacylglycerols in ozone-fumigated spinach leaves. Plant Physiol 94 (1990) 766-772
183. Benson A.A. The plant sulfolipid. Adv Lipid Res 1 (1963) 387-394
184. Benning C. Biosynthesis and function of the sulfolipid sulfoquinovosyl diacylglycerol. Annu Rev Plant Physiol Plant Mol Biol 49 (1998) 53-75
185. Harwood J.L. Sulfolipids. In: Stumpf P.K., and Conn E.E. (Eds). The biochemistry of plants. Lipids: structure and function vol. 4 (1980), Academic Press, New York 301-320
186. Mudd J.B., and Kleppinger-Sparace K.F. Sulfolipids. In: Stumpf P.K. (Ed). The biochemistry of plants. Lipids: structure and function vol. 9 (1987), Academic Press, New York 275-288
187. Harwood J.L., and Okanenko A.A. Sulphoquinovosyl diacylglycerol - the sulpholipid of higher plants. In: Abrol Y.P., and Ahmad A. (Eds). Sulphur in plants (2003), Kluwer, Dordrecht 189-219
188. Benning C. Questions remaining in sulfolipid biosynthesis: a historical perspective. Photosynth Res 92 (2007) 199-203
189. Haas R., Siebertz H.P., Wrage K., and Heinz E. Localization of sulfolipid labeling within cells and chloroplasts. Planta 148 (1980) 238-244
190. 14C]acetate in isolated intact spinach chloroplasts. Biochim Biophys Acta 879 (1986) 78-87
191. Heinz E., Schmidt H., Hoch M., Jung K.H., Binder H., and Schmidt R.R. Synthesis of different nucleoside 5′-diphospho-sulfoquinovoses and their use for studies on sulfolipid biosynthesis in chloroplasts. Eur J Biochem 184 (1989) 445-453
192. Seifert U., and Heinz E. Enzymatic characteristics of UDP-sulfoquinovose:diacylglycerol sulfoquinovosyltransferase from chloroplast envelopes. Bot Acta 105 (1992) 197-205
193. Pugh C.E., Roy A.B., Hawkes T., and Harwood J.L. A new pathway for the synthesis of the plant sulpholipid, sulphoquinovosyldiacylglycerol. Biochem J 309 (1995) 513-519
194. Essigmann B., Güler S., Narang R.A., Linke D., and Benning C. Phosphate availability affects the thylakoid lipid composition and the expression of SQD1, a gene required for sulfolipid biosynthesis in Arabidopsis thaliana. Proc Natl Acad Sci USA 95 (1998) 1950-1955
195. Mulichak A.M., Theisen M.J., Essigmann B., Benning C., and Garavito R.M. Crystal structure of SQD1, an enzyme involved in the biosynthesis of the plant sulfolipid headgroup donor UDP-sulfoquinovose. Proc Natl Acad Sci USA 96 (1999) 13097-13102
196. Sanda S., Leustek T., Theisen M.J., Garavito R.M., and Benning C. Recombinant Arabidopsis SQD1 converts UDP-glucose and sulfite to the sulfolipid head group precursor UDP-sulfoquinovose in vitro. J Biol Chem 276 (2001) 3941-3946
197. Yu B., Xu C., and Benning C. Arabidopsis disrupted in SQD2 encoding sulfolipid synthase is impaired in phosphate-limited growth. Proc Natl Acad Sci USA 99 (2002) 5732-5737
198. Shimojima M., and Benning C. Native uridine 5′-diphosphate-sulfoquinovose synthase, SQD1, from spinach purifies as a 250-kDa complex. Arch Biochem Biophys 413 (2003) 123-130
199. Shimojima M., Hoffmann-Benning S., Garavito R.M., and Benning C. Ferredoxin-dependent glutamate synthase moonlights in plant sulfolipid biosynthesis by forming a complex with SQD1. Arch Biochem Biophys 436 (2005) 206-214
200. Okazaki Y., Shimojima M., Sawada Y., Toyooka K., Narisawa T., Mochida K., et al. A chloroplastic UDP-glucose pyrophosphorylase from Arabidopsis is the committed enzyme for the first step of sulfolipid biosynthesis. Plant Cell 21 (2009) 892-909
201. Essigmann B., Hespenheide B.M., Kuhn L.A., and Benning C. Prediction of the active-site structure and NAD(+) binding in SQD1, a protein essential for sulfolipid biosynthesis in Arabidopsis. Arch Biochem Biophys 369 (1999) 30-41
202. Tietje C., and Heinz E. Uridine-diphospho-sulfoquinovose:diacylglycerol sulfoquinovosyltransferase activity is concentrated in the inner membrane of chloroplast envelopes. Planta 206 (1998) 72-78
203. Moore T.S. Phospholipid biosynthesis. Annu Rev Plant Physiol 33 (1982) 235-259
204. Andrews J., and Mudd J.B. Phosphatidylglycerol synthesis in pea chloroplasts: pathway and localization. Plant Physiol 79 (1986) 259-265
205. Müller F., and Frentzen M. Phosphatidylglycerophosphate synthases from Arabidopsis thaliana. FEBS Lett 509 (2001) 298-302
206. Xu C., Härtel H., Wada H., Hagio M., Yu B., Eakin C., et al. The pgp1 mutant locus of Arabidopsis encodes a phosphatidylglycerolphosphate synthase with impaired activity. Plant Physiol 129 (2002) 594-604
207. Babiychuk E., Müller F., Eubel H., Braun H.P., Frentzen M., and Kushnir S. Arabidopsis phosphatidylglycerophosphate synthase 1 is essential for chloroplast differentiation, but is dispensable for mitochondrial function. Plant J 33 (2003) 899-909
208. Tjellström H., Andersson M.X., Larsson K.E., and Sandelius A.S. Membrane phospholipids as a phosphate reserve: the dynamic nature of phospholipid-to-digalactosyl diacylglycerol exchange in higher plants. Plant Cell Environ 31 (2008) 1388-1398
209. Gao J, Ajjawi I, Manoli 3rd A, Sawin A, Xu C, Froehlich JE, et al. FATTY ACID DESATURASE 4 of Arabidopsis encodes a protein distinct from characterized fatty acid desaturases. Plant J 2009 [Epub ahead of print].
210. Schmidt H., and Heinz E. Desaturation of oleoyl groups in envelope membranes from spinach chloroplasts. Proc Natl Acad Sci USA 87 (1990) 9477-9480
211. Lawlor D.W. Photosynthesis: molecular, physiological and environmental processes. 2nd ed. (1993), Longmann Scientific and Technical, Harlow
212. Douce R., and Joyard J. Biosynthesis of thylakoid membrane lipids. In: Ort D.R., and Yocum C.F. (Eds). Advances in photosynthesis. Oxygenic photosynthesis: the light reactions vol. 4 (1996), Kluwer Academic Publishers, Dordrecht 69-101
213. Rawyler A., Meylan-Bettex M., and Siegenthaler P.A. (Galacto)lipid export from envelope to thylakoid membranes in intact chloroplasts. II. A general process with a key role for the envelope in the establishment of lipid asymmetry in thylakoid membranes. Biochim Biophys Acta 1233 (1995) 123-133
214. Carde J.P., Joyard J., and Douce R. Electron microscopic studies of envelope membranes from spinach plastids. Biol Cell 44 (1982) 315-324
215. Kroll D., Meierhoff K., Bechtold N., Kinoshita M., Westphal S., Vothknecht U.C., et al. VIPP1, a nuclear gene of Arabidopsis thaliana essential for thylakoid membrane formation. Proc Natl Acad Sci USA 98 (2001) 4238-4242
216. Liu C., Willmund F., Golecki J.R., Cacace S., Heß B., Markert C., et al. The chloroplast HSP70B-CDJ2-CGE1 chaperones catalyse assembly and disassembly of VIPP1 oligomers in Chlamydomonas. Plant J 50 (2007) 265-277
217. Wang Q., Sullivan R.W., Kight A., Henry R.L., Huang J., Jones A.M., et al. Deletion of the chloroplast-localized thylakoid formation1 gene product in Arabidopsis leads to deficient thylakoid formation and variegated leaves. Plant Physiol 136 (2004) 3594-3604
218. Huang J., Taylor J.P., Chen J.G., Uhrig J.F., Schnell D.J., Nakagawa T., et al. The plastid protein thylakoid formation1 and the plasma membrane G-protein GPA1 interact in a novel sugar-signaling mechanism in Arabidopsis. Plant Cell 18 (2006) 1226-1238
219. Andersson M.X., and Sandelius A.S. A chloroplast-localized vesicular transport system: a bio-informatics approach. BMC Genomics 5 (2004) 40
220. Blée E. Phytooxylipins and plant defense reactions. Prog Lipid Res 37 (1998) 33-72
221. Kachroo A., and Kachroo P. Fatty acid-derived signals in plant defense. Annu Rev Phytopath 47 (2009) 153-176
222. Ishiguro S., Kawai-Oda A., Ueda J., Nishida I., and Okada K. The DEFECTIVE IN ANTHER DEHISCIENCE gene encodes a novel phospholipase A1 catalyzing the initial step of jasmonic acid biosynthesis, which synchronizes pollen maturation, anther dehiscence, and flower opening in Arabidopsis. Plant Cell 13 (2001) 2191-2209
223. Feussner I., and Wasternack C. The lipoxygenase pathway. Annu Rev Plant Biol 53 (2002) 275-297
224. Schaller A, Stintzi A. Enzymes in jasmonate biosynthesis - structure, function, regulation. Phytochemistry 2009 [Epub ahead of print].
225. Blée E., and Joyard J. Envelope membranes from spinach chloroplasts are a site of metabolism of fatty acid hydroperoxides. Plant Physiol 110 (1996) 445-454
226. Froehlich J.E., Itoh A., and Howe G.A. Tomato allene oxide synthase and fatty acid hydroperoxide lyase, two cytochrome P450s involved in oxylipin metabolism, are targeted to different membranes of chloroplasts. Plant Physiol 125 (2001) 306-317
227. Austin II J.R., Frost E., Vidi P.A., Kessler F., and Staehelin L.A. Plastoglobules are lipoprotein subcompartments of the chloroplast that are permanently coupled to thylakoid membranes and contain biosynthetic enzymes. Plant Cell 18 (2006) 1693-1703
228. Joyard J, Ferro M, Masselon C, Seigneurin-Berny D, Salvi D, Garin J, et al. Chloroplast proteomics and the compartmentation of plastidial isoprenoid biosynthetic pathways. Mol Plant 2009 [in press]. doi:10.1093/mp/ssp088.
229. Stenzel I., Hause B., Miersch O., Kurz T., Maucher H., Weichert H., et al. Jasmonate biosynthesis and the allene oxide cyclase family of Arabidopsis thaliana. Plant Mol Biol 51 (2003) 895-911
230. Yokoyama M., Yamaguchi S., Inomata S., Komatsu K., Yoshida S., Ida T., et al. Stress-induced factor involved in flower formation of Lemna is an alpha-ketol derivative of linolenic acid. Plant Cell Physiol 41 (2000) 110-113
231. Lorenzo O., and Solano R. Molecular players regulating the jasmonate signalling network. Curr Opin Plant Biol 8 (2005) 532-540
232. Bell E., Creelman R.A., and Mullet J.E. A chloroplast lipoxygenase is required for wound-induced jasmonic acid accumulation in Arabidopsis. Proc Natl Acad Sci USA 12 92 (1995) 8675-8679
233. Park J.H., Halitschke R., Kim H.B., Baldwin I.T., Feldmann K.A., and Feyereisen R. A knock-out mutation in allene oxide synthase results in male sterility and defective wound signal transduction in Arabidopsis due to a block in jasmonic acid biosynthesis. Plant J 31 (2002) 1-12
234. Joyard J., and Stumpf P.K. Characterisation of an acyl-CoA thioesterase associated with the envelope of spinach chloroplasts. Plant Physiol 65 (1980) 1039-1043
235. Heinz E., Bertrams M., Joyard J., and Douce R. Demonstration of an acyltransferase activity in chloroplast envelopes. Z Pflanzenphysiol 87 (1978) 325-331
236. Andersson M.X., Hamberg M., Kourtchenko O., Brunnström A., McPhail K.L., Gerwick W.H., et al. Oxylipin profiling of the hypersensitive response in Arabidopsis thaliana. Formation of a novel oxo-phytodienoic acid-containing galactolipid, Arabidopside E. J Biol Chem 281 (2006) 31528-31537
237. Buseman C.M., Tamura P., Sparks A.A., Baughman E.J., Maatta S., Zhao J., et al. Wounding stimulates the accumulation of glycerolipids containing oxophytodienoic acid and dinor-oxophytodienoic acid in Arabidopsis leaves. Plant Physiol 142 (2006) 28-39
238. Andersson M.X., Goksör M., and Sandelius A.S. Optical manipulation reveals strong attracting forces at membrane contact sites between endoplasmic reticulum and chloroplasts. J Biol Chem 282 (2007) 1170-1174
239. Andersson M.X., Goksör M., and Sandelius A.S. Membrane contact sites: physical attachment between chloroplasts and endoplasmic reticulum revealed by optical manipulation. Plant Signal Behav 2 (2007) 185-187