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'Ioo_o.9. . . . 1-11 0 o .6: do. . . .. . .O‘. . o t. 3.0... .. or... to: , o..-o.o-.. . .i av?! .A. . . .. . u 5.. ..On. 7’... ...\ ‘O.l‘...‘\utu..~X-1ofl..' . 25 0’. . o i... )VI Q . ‘7. c.47'.‘0\§: Cl..-~\‘u 7.2 . .~...V.... .uo'v o....~¢6.:.0 . l'7 I» 1...! .1...‘ ' 3‘3 CJCQ 'f—_\ ‘J LIBRARY Michigan State University This is to certify that the dissertation entitled REGULATION OF LIPID METABOLISM IN RESPONSE TO ENVIRONMENTAL STRESS IN PLANTS AND ALGAE PhD. presented by Eric R. Moellering has been accepted towards fulfillment of the requirements for the degree in Biochemistry & Molecular Biology "iv/7% Major Professor’s Sighattfre g / /3( {D Date MSU is an Affirmative Action/Equal Opportunity Employer PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE 5/08 K:IProj/Aoc&Pres/ClRC/DateDue.indd REGULATION OF LIPID METABOLISM IN RESPONSE TO ENVIRONMENTAL STRESS IN PLANTS AND ALGAE By Eric R. Moellering A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Biochemistry and Molecular Biology 2010 ABSTRACT REGULATION OF LIPID METABOLISM IN RESPONSE TO ENVIRONMENTAL STRESS IN PLANTS AND ALGAE By Eric R. Moellering Glycerolipids are the major building blocks of biological membranes which provide semi-permeable boundaries for plant cells and the organelles within. Insight into the molecular and genetic factors required for lipid biosynthesis is crucial for understanding membrane processes as well as organelle biogenesis. The chloroplasts of eukaryotic plant cells are comprised of three membrane systems: the inner and outer envelope membranes, and thylakoids. The lipid composition is distinct in chloroplast membranes in that phosphorous—free galactoglycerolipids are the predominant class. The central genes and enzymes in galactolipid biosynthesis have been identified in Arabidopsis thaliana through combined molecular and genetic approaches. While a clear overall picture of galactolipid biosynthesis in plants has emerged, additional regulatory and metabolic factors remain to be discovered. A mutant suppressor screen was conducted in the Arabidopsis dgd] mutant deficient in the major biosynthetic pathway for the production of digalactosyldiacylglycerol. One mutant, dgdl suppressor 1 (dgsl) was identified and was found to cause a mutation in a mitochondrial protein of unknown function, DGS 1. Chapter 2 of this dissertation focuses on a detailed characterization of DGS 1. The dgsI-I mutation is semi-dominant and suppresses dng through the activation of (an) alternative biosynthetic pathway(s), likely through increased levels of reactive oxygen species present in dgsl—I . Importantly, analysis of a null allele, dgsI-Z, clearly revealed that effects on chloroplast localized galactolipid metabolism were only found in dgsI-I backgrounds, indicating that the wild-type DGS] protein is not directly involved in regulating galactolipid metabolism. However, dgs] -1 and other suppressors of dng were found to activate a poorly understood and distinct pathway for galactolipid biosynthesis via the enzyme galactolipidzgalactolipid galactosyltransferase (GGGT). Through a reverse genetics approach, the gene encoding GGGT was identified and found to be encoded by a previously described gene Sensitive T o Freezing 2 (SFR2). SFR2 encodes a glycosyl hydrolase family 1 protein required for freezing tolerance, and the work described in chapter 3 shows that SFR2 is sufficient for GGGT activity when expressed in heterologous form. The role for GGGT in fi'eezing tolerance is as a galactolipid remodeling enzyme that alters ratios of non-bilayer to bilayer forming lipids in the chloroplast envelope, which is hypothesized to stabilize this membrane during freeze induced dehydration. In a third project, the biosynthesis of triacylglycerols in the green alga Chlamydomonas reinhardtii during nitrogen deprived growth was investigated. TAGS accumulate during N-deprivation, and lipid droplets were isolated from cells under these conditions. Identification of the associated proteins (259 in total) revealed the existence of one predominant protein designated major lipid droplet protein (MLDP). MLDP is a protein with no known functional domains and its knockdown through RNA interference revealed a potential role for MLDP in lipid droplet morphology. In general, molecular factors involved in green alga are less characterized than their plant counterparts, and this work contributes initial insight into the cell biology of lipid droplets in green algal cells. ACKNOWLEDGEMENTS As I write this—the last section of my dissertation—thoughts and memories of all who have helped me along the way become overpowering. The past five years I have spent as a graduate student at Michigan State University have been the most challenging, and at the same time, most rewarding of my life. However, I must step back prior to my days at Michigan State, and acknowledge Dr. Raymond Chollet, now W.W. Marshall Family University Professor Emeritus at the University of Nebraska-Lincoln. My work as an undergraduate researcher in the Chollet lab solidified my passion for scientific inquiry, and provided an ideal combination of tutelage and challenge that few receive at this stage in their scientific career. I joined Dr. Christoph Benning’s lab at Michigan State in May, 2006, and while the time seems to have flown right by, it has truly been the most enjoyable of my life. From the beginning, I feel that Christoph showed trust in my abilities, and allowed me a great deal of freedom in finding my own way—all-the-while providing guidance to keep me on track. This is a mark of a truly great professor, and Christoph has been a mentor and friend to me, both professionally and personally, in too many ways to account for here. The work enviromnent he has created in his lab is a fertile ground for discovery, which one can only hope to find in the next step of their career, or one day replicate. With this in mind, I must acknowledge the Benning lab colleagues whom I’ve had the privilege to work with over the past five years. Numbering more than 30 over my stay at Michigan State, I’ll keep this general—mainly for fear of omission: you have all made work a joy to come to every day. The insight, debate, collaboration, jokes, quirks, iv frustration, and collective problem solving that we’ve all shared has been a blessing that is only rarely found at “work”. More broadly, I must acknowledge the institutions within Michigan State University that provided integral support during my graduate work. Firstly, the Department of Biochemistry and Molecular Biology, in which my degree is based, has offered nothing but premiere education to a graduate student. The Department of Energy Plant Research Laboratory—across the street from my BMB home base—is a truly unique feature of Michigan State (and a one of a kind worldwide institution), and I have been profoundly impacted as a graduate student in the PRL program. The above-affiliated faculty and staff, and the facilities maintained within the College of Natural Sciences have provided me with great opportunity and no restrictions in my research at Michigan State. In addition, the members of my guidance committee, Drs. Beronda Montgomery, Gregg Howe, Shelagh Ferguson-Miller, and John Ohlrogge have provided much appreciated support in my research. Funding provided by grants to Dr. Benning from the Department of Energy and the Air Force Office of Scientific Research were crucial in my doctoral research. I also want to thank all the fiiends I have gained while in Michigan. My time as a graduate student has been greatly enriched by spending our little free time together. Lastly, but certainly not least, I must express great love for my family—my parents and two brothers, for their love and support. TABLE OF CONTENTS List of Tables ..................................................................................................................... ix List of Figures ...................................................................................................................... x CHAPTER 1 Regulation of chloroplast glycerolipid biosynthesis in response to environmental stress...l Introduction .............................................................................................................. 2 A general overview fatty acid biosynthesis and glycerolipid assembly pathways in plants ........................................................................................................................ 3 Galactolipid biosynthesis during normal and nutrient limited growth conditions. . .............................................................................................................. 7 Galactolipid biosynthesis in non-photosynthetic tissues ....................................... 10 Galactolipid remodeling and turnover during various environmental stresses ...... 12 Production of cycIo-oxylipin-galactolipids during plant wounding and pathogen stresses ................................................................................................................... 13 Other routes for galactolipid biosynthesis in plants ............................................... l6 Biosynthesis of the chloroplast anionic glycerolipids SQDG and PG, during normal and stressed growth conditions .................................................................. l9 Perspectives ............................................................................................................ 22 References .............................................................................................................. 23 CHAPTER 2 Functional characterization of the DGS] protein in Arabidopsis thaliana ........................ 31 Abstract .................................................................................................................. 32 Introduction ............................................................................................................ 34 Materials and Methods ........................................................................................... 37 Plant materials and growth conditions ....................................................... 37 Bioinformatic analysis of DGS] ................................................................ 37 Generation of the DGS] antibody and irnmunoblot analysis .................... 38 Isolation of mitochondria and chloroplasts ................................................ 39 RNA analysis ............................................................................................. 40 Lipid analysis ............................................................................................. 40 Results .................................................................................................................... 41 Introduction of dgs1-2 does not suppress the dng growth phenotype ..... 4l DGSl is a mitochondrial membrane protein ............................................. 4] DGS] is present in a high molecular weight complex ............................... 46 Phosphate deprivation and dgsI-l allelism have additive effects on lipid metabolism ................................................................................................. 48 Alternative oxidase protein levels are reduced in the dgsI-I mutant allele .................................................................................................................... 53 The dgsl -1 mutant shows phenotypes of an AOXla-deficient mutant ..... 59 Discussion .............................................................................................................. 61 References .............................................................................................................. 64 vi CHAPTER 3 Identification of the gene encoding galactolipid2galactolipid galactosyltransferase and its role in freezing tolerance .................................................................................................. 69 Abstract .................................................................................................................. 70 Introduction ............................................................................................................ 71 Materials and Methods ........................................................................................... 72 Plant growth conditions ............................................................................. 73 Isolation of sfrZ T-DNA mutants and complementation of sfr2-3 with SFR2 cDNA ............................................................................................... 73 Lipid analysis ............................................................................................. 74 Heterologous expression of SFR2 in yeast ................................................ 75 Protein quantification and irnmunoblotting ............................................... 76 In vitro GGGT assay .................................................................................. 76 Results .................................................................................................................... 77 Development of a TLC-based screen to identify Arabidopsis mutants defective in GGGT activity ........................................................................ 77 T-DNA mutants in the SFR2 gene are defective in oligogalactolipid accumulation .............................................................................................. 79 The sfr2 mutants show defects in lipid remodeling during freezing stress .................................................................................................................... 79 Effect of various salts and osmolytes on oligogalactolipid and TAG accumulation following vacuum infiltration .............................................. 85 Genetic complementation of sfr2-3 by transgenic over-expression of the SFR2 cDNA ............................................................................................... 91 Heterologous expression of the SFR2 protein in yeast and analysis of in vitro GGGT activity ................................................................................... 91 Discussion .............................................................................................................. 95 A hypothesis for the role of GGGT in stabilizing chloroplast envelopes during freeze-induced cellular dehydration ............................................. 100 Evolution of glycosyl hydrolase family enzymes towards catalysis of transglycosidation: a proposed mechanism for GGGT activity ............... 103 Is the protective role of GGGT specific to freeze-induced dehydration? .................................................................................................................. 104 References ............................................................................................................ 108 CHAPTER 4 Literature Review: Molecular genetics of lipid metabolism in the green alga Chlamydomonas reinhardlii ............................................................................................ l 14 Abstract ................................................................................................................ 115 Abbreviations ....................................................................................................... l 16 Introduction .......................................................................................................... 1 17 Discussion ............................................................................................................ 118 Fatty acid synthesis and incorporation into glycerolipids ........................ l 18 Chloroplast membrane lipids ................................................................... 122 Extrachloroplastic membrane lipid metabolism ...................................... 127 vii Fatty acid desaturation ............................................................................. 137 Distinctions between ER-to-chloroplast lipid trafficking in plants and algae ......................................................................................................... l 36 Neutral glycerolipid metabolism .............................................................. 138 Perspectives .......................................................................................................... 141 References ............................................................................................................ 141 CHAPTER 5 Analysis of triacylglycerol accumulation in the green alga Chlamydomonas reinhardtii during nitrogen deprivation: Identification and characterization of an algal specific lipid droplet protein MLDP ...................................................................................................... 154 Abstract ................................................................................................................ 155 Introduction .......................................................................................................... l 56 Materials and Methods ......................................................................................... 158 Strains and growth conditions .................................................................. 158 Isolation of lipid droplets from N-limited C. reinhardrii ......................... 158 Mass spectrometric identification of lipid droplet proteins ..................... 159 Fatty acid and lipid analysis ..................................................................... 160 Construction of vectors and transformation of C. reinhardrii ................. 16] Quantitative Real-time RT-PCR .............................................................. 162 Fluorescent, confocal, and electron microscopy ...................................... 163 Bioinformatics .......................................................................................... 164 Results .................................................................................................................. 165 TAG accumulation following shift to N-limiting medium ...................... 165 Properties of lipid droplets ....................................................................... 168 Proteins identified in the lipid droplet fraction ........................................ 173 MLDP suppression affects lipid droplet size ........................................... 182 MLDP has orthologs only in green algae ................................................ 187 Discussion ............................................................................................................ 190 References ............................................................................................................ 1 95 CHAPTER 6 Conclusions and perspectives .......................................................................................... 199 DGSl is not directly involved in the regulation of galactolipid biosynthesis in plants. ................................................................................................................... 201 GGGT is encoded by the SFR2 gene in Arabidopsis and galactolipid remodeling in the chloroplast envelope is essential for freezing tolerance. ........................... 203 Triacylglycerol accumulation in Chlamydomonas reinhardtii: a role for MLDP in lipid droplet biogenisis? ...................................................................................... 207 References ............................................................................................................ 209 viii LIST OF TABLES Table 5.1. Total fatty acid content of whole cells compared to fatty acids bound in triacylglycerols isolated from cells which were grown in medium containing 10 mM NH;+ (+N), or cells switched to 0 mM NH4+ and grown for one day (-N) ..................... 167 Table 5.2. Fatty acid content of whole organelles (eyespot and lipid droplet) compared to fatty acids bound in triacylglycerols isolated from these organelles ............................... 172 Table 5.3. Complete list of proteins identified in the Chlamydomonas lipid droplet fraction ............................................................................................................................. 175 LIST OF FIGURES Images in this dissertation are presented in color. Figure 1.1. Glycerolipid assembly and trafficking in chloroplast membrane lipid biosynthesis .......................................................................................................................... 4 Figure 1.2 Chloroplast glycerolipid and fatty acid structures .............................................. 5 Figure 1.3 The four major chloroplast localized glycerolipid biosynthetic pathways ......... 8 Figure 1.4. C yclo—oxylipid-galactolipids (cGLs) produced during environmental stresses in Arabidopsis .................................................................................................................... 14 Figure 1.5. The GGGT galactolipid remodeling reaction .................................................. 17 Figure 2.1. Comparison of the dgs1-2 T-DNA null allele and the dgsI-l point mutant gain of function allele in wild type and dgdl backgrounds ............................................... 42 Figure 2.2. Sequence analysis of DGS] ............................................................................. 43 Figure 2.3. Localization of DGSl in mitochondria ........................................................... 45 Figure 2.4. Blue Native PAGE analysis of the DGS] complex ......................................... 47 Figure 2.5. Mitochondria of dgdl dgsl -1 homozygous mutant plants do not accumulate galactoglycerolipids ........................................................................................................... 49 Figure 2.6. Galactoglycerolipid content in the dgdl single homozygous mutant and the dng dgsI-I and dgd] dgsI-Z double homozygous mutants under normal and phosphate deprived growth conditions ................................................................................................ 51 Figure 2.7. Galactolipid Fatty acid profiles ...................................................................... 54 Figure 2.8. Alternative oxidase is reduced in dgsl-I mutant mitochondria ...................... 56 Figure 2.9. Analysis of AOXIa transcript levels in seedlings treated with Antimycin A... ............................................................................................................................................ 58 Figure 2.10. The dgsI -1 but not the dgs1-2 mutant displays a root growth phenotype ..... 60 Figure 3.1. Activation of GGGT activity in leaf tissue by vacuum infiltration of MgClz ............................................................................................................................................ 78 Figure 3.2. Genotypic confirmation of the sfr2 T-DNA mutant alleles used in this study. ................................................................................................................................ 80 Figure 3.3. Accumulation of oligogalactolipids in galactolipid biosynthetic mutants ...... 82 Figure 3.4. Freeze-induced galactolipid remodeling is not observed in sfr2 mutants ....... 83 Figure 3.5. Freeze-induced triacylglycerol accumulation .................................................. 86 Figure 3.6. ESl-MS/MS of triacylglycerols isolated from freeze-treated wild-type plants ............................................................................................................................................ 88 Figure 3.7. Salt and osmolyte treatments that activate TGDG and TAG accumulation in wild-type leaves ................................................................................................................. 89 Figure 3.8. Complementation of the sfr2-3 T-DNA mutant with SF R2 cDNA ................. 92 Figure 3.9. Heterologous expression of SFR2 in Saccharomyces cerevisae ..................... 94 Figure 3.10. Recombinant SFR2 in vitro activity from transgenic yeast ........................... 96 Figure 3.11. lH-NMR spectra of galactolipids from plant leaves and isolated from the in vitro GGGT assay .............................................................................................................. 98 Figure 3.12. Model of SF R2 activity and the effect of freezing on the chloroplast outer envelope membrane (oEv) and adjacent cell membranes (CM) in the sfi'2 mutant and wildtype (SFR2) ............................................................................................................... 102 Figure 3.13. A proposed reaction mechanism for GGGT ................................................ 105 Figure 3.14. Phylogenetic relationships of SFR2 orthologs ............................................ 107 Figure 4.1. Overview of glycerolipid biosynthesis in Chlamydomonas .......................... 120 Figure 4.2. Structures of PC and DGTS .......................................................................... 128 Figure 4.3. Overview of acyl-chain desaturation in (’hlamydomonas ............................. 132 Figure 5.1. Accumulation of triacylglycerols in lipid droplets during N-limitation ........ 166 Figure 5.2. Ultrastructure of cells during N-limitation .................................................... 169 Figure 5.3. Characteristics of isolated lipid droplets ....................................................... 170 Figure 5.4. Distribution of lipid droplet associated proteins into functional groups ....... 174 Figure 5.5. MLDP mRNA is induced following nitrogen deprivation ............................ 184 xi Figure 5.6. Generation of MLDP RNAi-mediated knockdown strains ........................... 185 Figure 5.7. MLDP RNAi knockdown lines exhibit a lipid droplet morphological phenotype ......................................................................................................................... 186 Figure 5.8. Multiple sequence alignment of MLDP with other green algal orthologs and hydropathicity plots of MLDP and other known lipid droplet binding proteins ............. 188 xii Chapter 1 Regulation of chloroplast glycerolipid biosynthesis in response to environmental stress Introduction Membranes provide boundaries for plant cells and the subcellular organelles within. Additionally, membranes are important sites for energy and signal transduction. Glycerolipids are major building blocks of biological membranes and membrane biogenesis requires exquisite regulation at the level of biosynthesis and extensive trafficking of glycerolipids. This results in subcellular membranes with distinct lipid compositions and homeostatic control of this composition is important for their specialized functions within the cell. Chloroplast membranes are an excellent example of this phenomenon, where non-phosphorous glycolipids are the predominant species. The chloroplast lipid biosynthetic pathways require enzymes that are localized at both the inner and outer envelope membranes of the chloroplast, and moreover, require trafficking of precursor lipid substrates from within the plastid, as well as glycerolipid backbones that are assembled in the endoplasmic reticulum. During normal growth and development chloroplast lipid composition remains remarkably constant in order to maintain an optimal environment for the thylakoid-localized photosynthetic machinery. However, under a number of environmental stress conditions, plants adapt by altering chloroplast lipid composition. This chapter will outline the current state of understanding of chloroplast lipid biosynthesis, with a focus on the factors required for modulating lipid responses in this dynamic organelle in response to environmental stress. A general overview fatty acid biosynthesis and glycerolipid assembly pathways in plants. De novo fatty acid biosynthesis takes place in the plastid in plant cells, resulting in the production of 1620- and 18:1 (carbons : double bonds) acyl chains linked to acyl carrier proteins (ACPs) (Ohlrogge and Browse, 1995, Ohlrogge et al., 1979). These fatty acids can be directly incorporated into glycerolipids in the plastid through the stepwise activity of acyl ACP dependent glycerol-3-phosphate acyltransferase (GPAT) and lysophosphatidic acid acyltransferase (LPAAT) enzymes to produce phosphatidic acid (PA) (Figure 1.1) (Ohlrogge and Browse, 1995). PA is used directly for the biosynthesis of phosphatidyl glycerol (PG), which is the only major phospholipid present in thylakoids (Ohlrogge and Browse, 1995, Xu et al., 2002, Xu eta]., 2006). PA can be further converted to diacylglycerol (DAG) by a PA phosphatase, providing a substrate for the biosynthesis of the chloroplast glycoglycerolipids mono- and di-galactosyldiacylglycerol (MGDG and DGDG, respectively) and sulfoquinovosyldiacylglycerol (SQDG) as firrther discussed below (Figure 1.2). Alternatively, fatty acids can be exported after their liberation fi'om ACP to the endoplasmic reticulum (ER) where they are converted to acyl CoA thioesters and assembled in to glycerolipids by direct incorporation into the phosphatidylcholine (PC) pool by acyl CoAzlysoPC acyltransferase or other enzymes (Bates er al., 2007), such as a set of ER-localized GPAT and LPAAT enzymes (Ohlrogge and Browse, 1995). The ER lipid biosynthetic pathway provides lipid precursors for the biosynthesis of extraplastidic membrane lipids including phosphatidylethanolamine (PE) and phosphatidylcholine (PC). Irnportantly, glycerolipid backbones assembled in the ER Cytoplasm/ER cap. E > L-PAtr' > PA_"* DAG ' Acyl-CoA s r PC at ‘ I l j“‘~+ PC El l .5 K,» DGDG Fi=A gimme °E" i G3P [2+ L-PA PA DAG I '5" IZl/v> l\ I\ I III Acyl-ACP PG 6318006 TE] IE] -J v v Thyfakoids _ Plastic! Figure 1.1. Glycerolipid assembly and trafficking in chloroplast membrane lipid biosynthesis. Acyl ACPs generated by the fatty acid synthase complex (FAS, step 1) are directly utilized in the chlorOplast by GPAT and LPAAT enzymes (steps 2, 3) to produce PA which is used in the biosynthesis of chloroplast PG (step 7, multiple enzymes). Fatty acids hydrolyzed fiom Acyl ACPs in the plastid are exported to the cytosol and esterified in acyl CoAs, which serve as substrates for extraplastidic glycerolipid assembly through ER—localized GPAT and LPAAT (steps 9, 10) or through acyl editing of the PC pool (Step 13, multiple enzymes likely). Biosynthesis of extraplastidic phospholipids includes the major lipid, PC (via steps 11, 12), which provides the precursor backbone for glycerolipid equivalents that are imported from the ER into the chloroplast (step 14) and enters the chloroplast DAG pool which is also contributed to by the plastid glycerolipid assembly pathway via the dephosphorylation of PA by phosphatidic acid phosphatase (step 4). This DAG pool is used for the biosynthesis of the major galactolipids MGDG and DGDG (steps 5 and 6, respectively) and the sulfolipid SQDG (step 8, multiple enzymes). The glycolipids and PG are trafficked to the thylakoid (step 15) through mechanisms which are incompletely understood. Abbreviations: DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; FFA, free fatty acids; G3P, glycerol-3-phosphate; IBM, inner envelope membrane; L-PA, lysophosphatidic acid; MGDG, monogalactosyldiacylglycerol; OEM, outer envelope membrane; PA, phosphatidic acid; PC, phosphatidylcholine; PG, phosphatidylglycerol; SQDG, sulfquinovosyldiacylgycerol. 4 Figure 1.2. Chloroplast glycerolipid and fatty acid structures. (A) Structures for the galactolipids MGDG and DGDG, the phospholipids PC and PG, and the sulfolipid SQDG; abbreviations are as in Figure 1.]. Each glycerolipid has a distinct fatty acid composition esterified to the sn-1 and sn-2 position of the glycerolipid backbone; and the most common fatty acid species for A. thaliana (several others have been observed) is indicated by the R1 (for sn-I) and R2 (for sn-2) acyl group species indicated as letters A-F in parentheses. The identities of A-F are shown in (B). (B) Common fatty acid acyl groups of glycerolipids. The common names and abbreviation for the corresponding fatty acids are indicated as is the R-group identifier used in (A). In the drawn structures, the R- group linkage site is circled, and numbers above carbonzcarbon double bonds indicate the position counting from the carboxyl end. HO OH 0 \/L O- o H DGDG W 0 0’K P, R2 (A,B) HO 0 g/ 0 HO OH MGDG (DE/R1 (A'E) O O CH2303' SQDG Q o m am «a % H0 R2 (A E) OH 0 Trivial name R-group and abbreviation identifier Structure palmitic acid (16:0) AIS-trans hexadecenoic acid (16:1A3!) oleic acid (18:1) Iinoleic acid (18:2) a-Iinolenic acid (18:3) ‘nrnUOm> palmitolenic acid (16:3) pathway are trafficked to the chloroplast, feeding into the DAG pool used for the synthesis of the above-mentioned chloroplast glycolipids (Figure 1.1). Evidence for this bifurcation in glycerolipid assembly came first from substrate radiotracer studies (Roughan et al., 1980, Roughan and Slack, 1982). The “two pathway” hypothesis was later confirmed, when mutants in the plastidial GPAT and the ER and plastidial desaturases were identified in Arabidopsis (Browse and Somerville, 199], Kunst er al., 1988). Due to differences in substrate specificities of the plastid and ER-localized lipid assembly enzymes, glycerolipids derived from either pathway can be distinguished by the length of the acyl chain esterified to the sn-2 position—where plastid derived lipids contain 16-carbon acyl chains as opposed to the 18-carbon chains found in ER-derived lipids. The relative contributions of the plastid and ER-localized glycerolipid assembly pathways in chloroplast glycoglycerolipid biosynthesis varies among species—where in spinach and Arabidopsis for example, the plastidial pathway is significant; conu'astingly in many monocots and in pea, the ER pathway is the sole contributor (Mongrand el al., 1998). Though not a focus of this chapter, factors required for ER-to-chloroplast lipid trafficking have recently been identified (Benning, 2009). Galactolipid biosynthesis in leaves during normal and nutrient-limited growth conditions. The galactolipids MGDG and DGDG are the predominant glycerolipid species in chloroplasts (Dormann and Benning, 2002). The bulk of galactolipid biosynthesis is carried out by two UDP-galactose (UDP-Gal) dependent enzymes, MGDl and DGD] (see Figure 1.3) (Benning and Ohta, 2005). MGD] is localized to the inner chloroplast UTP PPi so3 G1P¥> UDP-GIG UDP-SQ UDP Pi PA DAG SQDG crp \ 0034/5 / UDP-Gal PPi/ i \ UDP COP-DAG i G3P \ MGDG K UDP-Gal CMP/ \ v UDP PGP y DGDG / V PG Figure 1.3. The four major chloroplast localized glycerolipid biosynthetic pathways. Substrates, enzymes and steps required for the biosynthesis of the four major chloroplast glycerolipids MGDG, DGDG, SQDG and PG. Enzymes are indicated in round-edged boxes, and full names for the abbreviations are: CDS, CDP-DAG synthase; DGD], DGDG synthase; MGDl, MGDG synthase; PAP, PA phosphatase; PGP], PGP synthase; PGPase, PGP phosphatase; SQDl, UDP-SQ synthase; SQD2, sulfolipid synthase; UGP3, UDP-Glc pyr0phosphorylase 3. Abbreviations for molecules are as in Figure 1.1, or: CDP-DAG, citidinediphosphate—diacylglycerol; CTP, citidinetriphosphate; GlP, glucose- l-phosphate; Pi, phosphate; PPi, pyrophosphate; 803', sulfite; UDP, uridinediphosphate, UDP-Gal, UDP-galactose; UDP-Glc, UDP—glucose; UDP-SQ, UDP-sulfoquinovose; UTP, uridinetriphosphate. envelope (Awai et al., 2006) where it converts DAG into MGDG. This MGDG can either be directly transported to the thylakoid membranes, or to the outer chloroplast envelope, where DGDl acts to convert MGDG to DGDG (Froehlich er al., 2001). A series of studies on Arabidopsis mutants in this pathway have revealed the importance of galactolipid biosynthesis during normal plant growth and development A leaky allele of mng was shown to have defects in chloroplast biogenesis (Jarvis e101,, 2000), and a more recently described null allele was found to be embryo or seedling—lethal (Kobayashi et al., 2007). A null allele for dng identified through forward genetics showed a severe dwarf growth phenotype, but ~10 % of wild-type DGDG levels remained (Diirmann er al., 1995, Dr'irmann er al., 1999). This latter observation, along with DGDG accumulation in the dgdl mutant during phosphate deprivation (Hartel er al. , 2000), indicated the presence of additional pathways for DGDG biosynthesis. Indeed, a reverse genetics approach identified genes encoding a paralog of DGDI, DGD2 (Kelly and Dormann, 2002). After initial identification of the MGDG synthetase purified from cucumber (Shimojima er al., 1997), its ortholog, MGDI, and two paralogs of MGDI, MGD2,3 were also identified in the Arabidopsis genome (Awai er al., 2001). MGD2,3 and DGD2 were shown to be coordinately regulated at the transcriptional level, and highly expressed under phosphate-limiting growth conditions. The MGD2,3 and DGD2 proteins are localized to the chloroplast outer envelope, and comprise an alternative pathway for galactolipid biosynthesis that produces DGDG that is trafficked to extraplastidic membranes (e. g. the tonoplast, mitochondria, and plasma membrane) during phosphate deprivation stress in a mechanism that conserves phosphate through the replacement of phospholipids (Hartel et al., 2000, Jouhet et al., 2004, Jouhet er al., 2007, Tjellstrom et al., 2010). In a detailed study of plasma membranes in phosphate limited oat (Avena satrva), replacement of phospholipids with DGDG was found to be restricted to the cytosolic leaflet (Tjellstrom et al., 2010). More recently, nitrogen (N) or magnesium deprivation was shown to alter galactolipid composition in Arabidopsis leaves, where MGDG was found to decrease, with lesser increases in both DGDG and phospholipids (Gaude et al., 2007). Analysis of transcript levels for the above-described galactolipid biosynthetic genes revealed increased expression of both DGDI and DGD2, but little or no change in the MGD1/2/3 genes. Fatty acid phytyl esters and triacylglycerols were found to accumulate in response to N-deprivation, and more detailed analysis revealed an increase in the amount of 16:3 (carbons : double bonds) fatty acid esterified to phytol. As 16:3 is predominately esterified to MGDG in Arabidopsis during normal growth (Browse 2:01., 1989), these results suggested that during N-deprivation, MGDG is being turned over and the fatty acids are converted in part to fatty acid phytyl esters (FAPEs) (Gaude e101,, 2007). However, the identities of potential lipase and/or transferase enzymes involved in the formation of FAPEs and their role during N-limitation in plants remain unknown. Galactolipid biosynthesis in non-photosynthetic tissues. Not surprisingly, the role of the phosphate-regulated MGD2/3-DGD2 dependent DGDG biosynthesis pathway is not restricted to photosynthetic tissues. As in leaves, MGD2/3 and DGD2 are activated in root tissue in Arabidopsis during phosphate deprivation (Andersson er al., 2003, Benning and Ohta, 2005, Kelly and Dormann, 2002, Kobayashi et al., 2004, Kobayashi et al., 2006, Lai et al., 2007). Additionally, DGDG 10 has been found to accumulate in phosphate-deprived roots from a number of plant species in addition to Arabidopsis, including Phaseolus vulgaris (bean) (Hartel er a1. , 2000, Russo er al., 2007). DGDG also accumulates in peribacteroid membranes (PBMs) in the legume species soybean and Lotus; the FEM is of plant origin, but surrounds the so- called “symbiosome” organelle containing nitrogen fixing bacterial species within the root nodule cells (Gaude er al., 2004). The DGDl/2 orthologs in soybean were shown to exhibit nodule-enhanced expression (compared to roots), and in situ hybridization indicated the DGDl/Z mRN As to be enriched in the infected cells of the nodule (Gaude er a1. , 2004). A role for galactolipid biosynthesis during pollen tube growth has also emerged (Kobayashi er al., 2004, Nakamura et al., 2009a). Transgenic Arabidopsis lines containing promoterzGUS fusion constructs for MGD1,2,3 indicated the expression of all three genes in anthers, whereas only MGD2 and MGD3 were expressed during pollen tube growth (Kobayashi etal., 2004). Analyses of lipid content in Lilium Iongzflorum revealed accumulation of MGDG and DGDG afier pollen tube elongation (Nakamura et al., 2009a). Given the rapid biosynthesis of membranes required for pollen tube elongation, a role for the alternative galactolipid biosynthetic pathway in providing extraplastidic DGDG for this process was hypothesized. However, whether or not mutants in the alternative galactolipid biosynthetic pathway are defective in pollen tube growth is currently not known, but major defects are unlikely as null mutants in these genes have been studied. 11 Galactolipid remodeling and turnover during various environmental stresses. Although most insight has been gained on the synthesis of galactolipids in plants, both chilling (Kaniuga and Gemel, 1984) and drought (N avari-lzzo and Rascio, I999, Torres-Franklin er al., 2007) stress conditions and senescence (Kaup et al., 2002) are associated with catabolism and/or remodeling of these predominant chloroplast lipids. In response to low temperature, chilling resistant plants generally show increased fatty acid desaturation levels (Iba, 2002), but these stress-induced changes are not specific to galactolipids or chloroplast glycerolipids in general, and as such are not further discussed here. In response to severe water deficit, the MGDGzDGDG ratio decreases markedly in Arabidopsis leaves, and transcript levels of MGDI were found to decrease during severe drought, while DGDI mRNA levels did not (Gigon et al., 2004). Decreases in the MGDGzDGDG ratio during water deficit and desiccation have been observed in a number of different plant species (Navari-lzzo and Rascio, 1999), suggesting a conserved but incompletely understood mechanism for chloroplast lipid remodeling during this stress. A role for galactolipid remodeling during acquired thermotolerance was revealed in Arabidopsis, when isolated thennosensitive mutants were mapped and identified as weak alleles of DGDI (dng-Z and dgd1-3) (Chen et al., 2006). The dng-Z and -3 alleles showed only ~40-50 % decreases in steady state DGDG levels and no apparent growth defects, and upon exposure to 38° C, did not show the increased DGDG content and thus altered MGDGzDGDG ratio observed in wild-type plants during heat stress (Chen et al., 2006). Similar changes have also been observed during exposure of plants to excess copper or cadmium (Chaffai et al., 2007, Jemal et al., 2000). Whether or not additional enzymes are involved in stress-induced galactolipid changes remains unclear. 12 In some cases, the changes in lipid composition may be due to changes in transcriptional or metabolic regulation of the known galactolipid biosynthetic pathways, or due to increased oxidation-dependent turnover rates that are an indirect consequence of some stresses (e.g. excess copper). Recently, a candidate a-galactosidase for the conversion of DGDl/Z derived DGDG to MGDG has been identified in 0ryza sativa (rice) (Lee et al., 2009, Lee el al., 2004). The encoding gene, OsAkaGal, produced a functional protein when expressed in E. coli, and a null allele in rice showed a delayed senescence phenotype (Lee et al., 2009). However, lipid composition was not reported for wild-type plants or the osakagal mutant and the in viva role for this enzyme in galactolipid metabolism remains to be established. Production of cyclo—oxylipin-galactolipids during plant wounding and pathogen stresses. MGDG and DGDG are rich in tri-enoic fatty acids (16:3 and 18:3 in Arabidopsis), and during stresses such as mechanical wounding (Buseman el al., 2006) or in response to induced accumulation of the Pseudomonas syn’ngae avirulence protein, Achme , in planta (Andersson et al., 2006), these galactolipid esterified fatty acids are converted to oxylipins, resulting in the increased formation of what are collectively referred to as cyclo-oxylipin-galactolipids (cGLs, also referred to as Arabidopsides A-G, see Figure 1.4) (Bottcher and Pollmann, 2009). cGL formation currently appears to be restricted to Arabidopsis and other related Brassicaceae (Bottcher and Pollmann, 2009). The esterified oxylipin species produced are lZ-oxo-phytodienoic acid (OPDA) or dinor- (dn)-OPDA l3 A B HO R3 anPDA OPDA 0 R2 H 0 HO 0” MGDG R1 HO | | O H HO 0 0.. $ ; R2 0 ° DGDG c R‘ . MGDG or cGL s I R R R pec es DGDG 1 2 3 MGDG—O M OPDA 16:3/18:3 -OH ArabidopsideA M OPDA OPDA -0H Arabidopside B M OPDA anPDA ‘0“ Arabidopside c o OPDA anPDA '3: Arabidopside o o OPDA OPDA N/A Arabidopside E M OPDA anPDA OPDA Arabidopside F M 18:3 OPDA .OH Arabidopside G M OPDA OPDA OPDA Figure 1.4. Cyclo—oxylipin-galactolipids (cGLs) produced during environmental stresses in Arabidopsis. (A) structures of (dinor)oxo-phydodienoic acids). (B) Galactolipids MGDG and DGDG with possible sites of (dn)OPDA esterification indicated by R], R2, and R3. (C) A legend to the diverse cGL species identified in Arabidopsis, including MGDG-0 and Arabidopsides A-G. The derived galactolipid is indicated (M or D for MGDG and DGDG, respectively) and the positions of fatty acids or their oxidized (dn)OPDA derivatives are indicated for depictied in R1, R2, and R3 (B). 14 from 18:3 and 16:3 fatty acids respectively by the same set of enzymes: A lipoxygenase (LOX) catalyzes the formation of hydroperoxy- fatty acids that are further converted to anPDA and OPDA by the stepwise activities of allene oxide synthase (A08) and allene oxide cyclase (AOC) (Howe and Schilmiller, 2002, Schaller and Stintzi, 2009). OPDA and anPDA serve as precursors for the biosynthesis of jasmonates, a class of plant hormones with diverse fimctions in the regulation of abiotic and pathogen stress responses as well as growth and development (Browse, 2009, Howe and Schilmiller, 2002, Schaller and Stintzi, 2009). Intriguingly, the levels of OPDAs esterified in cGLs is far higher than the corresponding free acids (e.g. ~100 fold in Aerme expressing plants (Andersson et al., 2006)), leading to the hypothesis that cGLs serve as a stored form of OPDAs that adds an additional level of regulating substrate levels feeding into the jasmonate biosynthetic pathway (Buseman et al., 2006). While evidence for direct biosynthesis of cGLs on galactolipid bound fatty acids is evident in the species formed, the identification of two new cGLs, Arabidopsides E and G (Figure l.4C), clearly indicate that further modification via transacylation is involved in their biosynthesis. Arabidopsides E and G are tri-oxylipin containing MGDG derivatives that, in addition to the OPDAs esterified to the sn-1 and -2 positions of the glycerol backbone found in other cGLs, contain a third OPDA esterified to the 6’ carbon of the galactosyl moiety. Arabidopside E (containing anPDA esterified to the sn-2 position of the glycerol backbone) was shown to inhibit grth of P. syringae, suggesting that some cGLs may play a direct role in pathogen defense (Andersson et al., 2006). While the acyltransferase involved in the biosynthesis of Arabidopsides E and G remains unknown, 15 it was hypothesized to be the same or similar to the unknown enzyme responsible for the synthesis of acyl-MGDG first identified over 50 years ago (Heinz, l967a, Heinz, l967b). Other routes for galactolipid biosynthesis in plants. A third and distinct pathway for galactolipid biosynthesis occurs through the activity of a galactolipidzgalactolipid galactosyltransferase (GGGT) enzyme (Figure 1.5) (Benning and Ohta, 2005). GGGT was first described at the biochemical level over 40 years ago (van Besouw and Winterrnans, 1978), but its encoding gene(s) and role in plants has remained enigmatic (Benning and Ohta, 2005, Kelly er al., 2003). Unlike the above-described MGDl/2/3 and DGDl/2 galactolipid synthases, GGGT is UDP-Gal independent (Heemskerk er al., 1983, Heemskerk et al., 1990, van Besouw and Wintermans, 1978), and instead uses two MGDG molecules as substrate to catalyze the formation of DGDG and DAG via transgalactosylation (Figure l.5A). The DGDG produced by GGGT is in the all-B conformation with respect to the glycosidic linkages (BBDGDG), which is distinct from the BaDGDG produced by DGDl/2 (Figure l.5B~C). Furthermore, through processive activity, GGGT catalyzes the formation of the oligogalactolipids tri- and tetra-galactosyldiacylglycerol (TGDG and TeDG, respectively) (van Besouw and Winterrnans, 1978). Due to the high activity of GGGT and lack of a UDP—Gal dependent effect on DGDG formation from in vitro biochemical studies in different species, GGGT was hypothesized to be the major contributor to DGDG biosynthesis in vivo (Heemskerk er al., 1983, Heemskerk et al., 1990, van Besouw and Wintermans, 1978). 16 0R2 Figure 1.5. The GGGT galactolipid remodeling reaction. (A) Three catalytic cycles of processive GGGT activity are shown, with the circled number indicating the order of reactions. The galactolipid products, BBDGDG, BBBTGDG, and BBflBTeDG are produced along with three DAG molecules. Other GGGT catalyzed transferases reactions are possible (e.g. transfer of a galactosyl moiety between two BBDGDG molecules). The difference between the structures of BnDGDG produced by DGDl/2 biosynthetic pathways (B) and BBDGDG produced by GGGT (C) are shown. The B or (1 letters for the galactolipid species indicate the anomeric configuration of the l-6 glycosidic linkages present in each molecule, beginning with the one most distal to the glycerol backbone moiety. For simplicity, the hydroxyl groups of the galactosyl head group moieties are drawn as sticks, and the galactosyl protons attached to the l’ C anomeric centers are indicated as H1’ and H3”. As described above, molecular genetics revealed a major role played by DGDl (Dormann e101,, 1999), and further analysis of the dgd] dgd2 double mutant revealed it to be devoid of detectable levels of DGDG (Kelly etal., 2003), whereas GGGT activity in isolated chloroplasts of dgdl dgd2 remained indistinguishable from wild-type. Indeed, GGGT was found to be activated due to osmotic shock arising from certain steps of intact chloroplast or envelope membrane isolation (Dome er al., 1982, Dome etal., 1985), suggesting a possible role for GGGT in response to water deficit. GGGT was also found to be activated in a series of studies on the effect of ozone (03) treatment on chloroplast lipid metabolism (Sakaki er al., l990a, Sakaki er al., 199%, Sakaki et al., l990c). O3 fumigated spinach leaves accumulated oligogalactolipids as well as triacyl glycerol (TAG) (Sakaki et al., 199%, Sakaki er al., l990c), and molecular analysis revealed that DAG derived from GGGT activity was being utilized as a substrate for TAG biosynthesis. The significance of these galactolipid remodeling events during ozone stress remained unclear. In vitro studies of GGGT enzyme activity indicated its activation by divalent cations (e.g. Mg”, Ca”, and Ba2+) as well as free fatty acids added to the assay (Heemskerk er al., 1983, Sakaki er al., l990a). Most recently GGGT has been found to be constitutively activated in vivo in the tgd ER-to—chloroplast lipid trafficking mutants (Awai et al., 2006, Benning, 2009, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008). Originally isolated as genetic suppressors of dgdl (Xu er al., 2003), the rgd1/2/3/4 mutants accumulate oligogalactolipids in shoot tissues, although the molecular/biochemical etiology behind the activation of GGGT in these mutants is currently unknown. During research conducted in the preparation of this dissertation, the gene encoding GGGT was identified in Arabidopsis and is further described in Chapter 3. 18 TGDG and/or TeDG are constitutively produced in certain tissues of certain plant species including rice bran, Adzuki bean, potato tubers, and pumpkin fruit (reviewed in (Holzl and D'o'rmann, 2007, Kelly and Dormann, 2004). However, the analysis of the anomeric configuration of the glycosidic linkagese for these constitutively produced oligogalactolipids (Fujino and Miyazawa, 1979, Galliard, 1969, Kojima et al., 1990) indicates biosynthetic pathways that are likely to be independent of the all B- configuration oligogalactolipids produced by GGGT in Arabidopsis (Xu et a]. , 2003)— possibly involving multiple enzymes. To date, no enzyme known to produce oligogalactolipids in plants has been characterized at the molecular/genetic level. Biosynthesis of the chloroplast anionic glycerolipids SQDG and PG during normal and stressed growth conditions. Although the head group moieties of the sulfolipid, SQDG, and phospholipid, PG, may appear to share little structural similarity, these lipids have similar roles in biological membranes due to the negative charge that they carry (see Figure 1.2). These anionic lipids are important for maintaining a negative charge density that is important for the proper function of the photosynthetic machinery (Wada and Murata, 2007). SQDG biosynthesis begins with the production of UDP-SQ by SQDl (in Arabidopsis) using UDP-Glc and sulfite as substrates (Figure 1.3). SQDG is then formed by SQD2, which transfers the sulfoquinovose moiety to DAG. This pathway was first elucidated in studies on Rhodobacter sphaeroides (Benning and Somerville, 1992a, Benning and Somerville, 1992b), and through analysis of Rhodobacrer mutants, SQDG biosynthesis was shown to be of conditional importance during phosphate deprivation l9 (Benning er al., 1993). An Arabidopsis sqd2 mutant lacking SQDG showed a similar conditional importance of sulfolipid in higher plants, where no growth defects were observed until squ was presented with phosphate-deprived conditions (Yu et al., 2002). The SQD1/2 genes show up-regulation during phosphate deprivation, and under these conditions, the SQDG levels increase with a concomitant decrease in PG, suggesting a homeostatic mechanism to maintain a negative charge balance under varying environmental conditions (Yu et al., 2002). The conditional importance of SQDG may not be universal among organisms which produce this lipid: a null mutant in Synechocystis PCC 6803 squ gene (an ortholog of Arabidopsis SQDI) is a sulfolipid auxotroph (Aoki et a1. , 2004). Co-expression of Arabidopsis SQDI and SQD2 genes heterologously in E. coli indicated that the products of these two genes are sufficient for the production of SQDG in plants (Y 11 et al., 2002). Recently, a third gene required for SQDG biosynthesis in planta has been identified Utilizing transcriptome co-expression data in Arabidopsis, Okazaki and colleagues identified a candidate UDP-Glc pyr0phosphorylase encoding gene (UGP3) that showed strong correlation of expression with known sulfolipid biosynthetic genes (Okazaki er al., 2009). Null mutants in UGP3 lacked SQDG, and a UGP-GFP fusion protein was targeted to the chloroplast (Okazaki er al., 2009), providing convincing evidence that UGP3 is a stroma-localized enzyme required for the synthesis of the UDP-Glc substrate for SQDI. PG is the only major phospholipid synthesized in the chloroplast, and as alluded to above, its biosynthetic pathway is distinct from that of the chloroplast glycolipids. As seen in Figure 1.3, PG biosynthesis begins with the production of CDP-DAG by the CD8 20 enzyme. Recent molecular/genetic studies in Arabidopsis have identified 5 candidate CDS encoding genes, and revealed that the CDS4/5 isogenes encode plastidial isozymes with overlapping functions in PG biosynthesis (Haselier er al., 2010). While no phenotypes were observed for cds4 or cds5 single mutants, isolation of the corresponding cds4 cds5 double mutant revealed severe growth phenotypes including non-viability on soil, and pale yellow leaves when grown on agar medium supplemented with sucrose, while showing a ~60 % decrease in total leaf PG when compared to wild-type (Haselier er al., 2010). CDP-DAG produced in the chloroplast is further converted to phosphatidylglycerolphosphate by PGP] (Figure 1.3) (Wada and Murata, 2007). The py] mutants in Arabidopsis show similar PG deficiencies and phenotypes as described for the cds4 cds5 double mutant (Babiychuk etal., 2003, Hagio er al., 2002, Xu et al., 2002). The last step in chloroplast PG biosynthesis is the dephosphorylation of PGP, which is catalyzed by a PGP phosphatase that remains unidentified in plants at the molecular/genetic level. The gene encoding PGP phosphatase have remained unknown in eukaryotes until only recently, when a mitochondrial protein required for cardiolipin biosynthesis in Saccharomyces cerevisae, Gep4, was shown to function as a PGP phosphatase (Osman et al., 2010). Intriguingly, a distant chloroplast targeted (predicted) ortholog of Gep4 is encoded in the Arabidopsis genome (by the gene At3G58830)—but its possible role in PG biosynthesis is not known. Modulation of PG levels during phosphate-limiting stress occurs (Yu et al., 2002), but it is clear from the PG deficient mutants that PG cannot be functionally replaced by SQDG in plants, where as the opposite appears to be the case during normal growth and development. 21 Perspectives Plants have evolved myriad responses to cope with changes in their environment, and the modulation of lipid metabolism during stress has emerged as a prominent factor for a number of different responses. While this chapter focused on changes in chloroplast lipid biosynthesis, responses of extraplastidic lipid metabolism during stress at least equal that within the chloroplast (Jouhet et al., 2007, Li et al., 2008, Welti etal., 2002). Indeed, these processes are not mutually exclusive as there is dynamic exchange of lipids between the chloroplast and extraplastidic membranes through extensive lipid remodeling and trafficking pathways during normal and stressed growth conditions (Benning, 2009, Cnrz-Ramirez et al., 2006, Gaude etal., 2008, Li et al., 2006, Nakamura e101,, 2009b). Since the release of the Arabidopsis genome nearly] 0 years ago (The Arabidopsis Genome Initiative, 2001), great progress has been made in understanding lipid biosynthetic pathways in plants (Wallis and Browse, 2010). However, several questions remain. Among these are: what are the regulatory and metabolic pathways required for lipid responses to environmental stress? Undoubtedly, the increasingly powerful “omics” approaches are going to play an expanding role in elucidating the many unknown facets in plant lipid metabolism (Okazaki et al., 2009). Additionally, the expansion of molecular tools in other model plants, e. g. 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B., and Benning, C. (2003) A permease-like protein involved in ER to thylakoid lipid transfer in Arabidopsis. EMBO J. 22, 2370- 2379. Xu, C., Harte], H., Wada, H., Hagio, M., Yu, B., Eakin, C., and Benning, C. (2002) The pg] locus of Arabidopsis encodes a phosphatidyl glycerol synthase with impaired activity. Plant Physiol. 129, 594-604. Yu, B., Xu, C., and Benning, C. (2002) Arabidopsis disrupted in SQD2 encoding sulfolipid synthase is impaired in phosphate-limited growth. Proc. Natl. Acad. Sci. U. S. A. 99, 5732-5737. 30 Chapter 2 Functional characterization of DGSI in Arabidopsis thaliana1 I This work has been published in: (i) Xu, C., Moellering, E.R., Fan, J ., and C. Benning. (2008) Mutation of a mitochondrial outer membrane protein affects chloroplast lipid biosynthesis. Plant J. 542163-175, and (ii) Moellering, ER. and C. Benning. (2010) Phosphate regulation of lipid biosynthesis in Arabidopsis is independent of the mitochondrial outer membrane DGSl complex. Plant Physiol 152: 1951-1959. Experiments for the data shown in Figure 2.3, panels A-B, were conducted by Changcheng Xu, as was the production of the antigenic polypeptide used in generating the DGSl antibody. 31 Abstract Galactolipid biosynthesis in plants is carried out by UDP-galactose utilizing enzymes that are localized to the chloroplast envelopes. In Arabidopsis, the bulk of digalactosyldiacylglycerol (DGDG) is produced by DGDI during normal grth conditions. A paralog of DGDl , DGD2, is induced during phosphate deprivation in a mechanism that replaces phospholipids with DGDG in extraplastidic membranes. In order to identify the largely unknown factors that regulate the DGDI-independent pathway, and possibly lipid homeostasis in general, a screen for suppressors of the galactolipid deficient dgd] mutant was conducted. One mutant, dgsI-l (dgdl suppressor 1-1), was identified with partially restored DGDG content. Initial analyses revealed the DGSI protein to be present in the mitochondria, and that the dgsI-l suppressor phenotype was semi-dominant. Moreover, dgsI-I plants were found to accumulate H202, which when applied to dgd] plants, was sufficient to activate dng-independent DGDG biosynthesis. Here, continued analysis of the gain-of-function dgsl -1 mutant was accompanied by analysis of a null T-DNA allele, dgsl-Z. The DGSI protein was found to be part of a large protein complex present in the outer mitochondrial membrane. The dgsl-I allele causes the loss of the mitochondrial alternative oxidase (AOX) protein which could possibly explain the accumulation of H202 in this background. The loss-of- function dgsl-2 allele had no observable effect on plant growth, AOX, or lipid composition. Moreover, accumulation of DGDG in phosphate-deprived dgdl dgsI -1 double mutants indicates that the dgsI -1-dependent DGDG accumulation is additive to the phosphate responsive pathway. These results indicate that DGSI is not directly 32 involved in the regulation of lipid metabolism in chloroplasts and mitochondria, and leaves the molecular function of this protein unresolved. 33 Introduction In plant cells, the membranes that provide boundaries for the various subcellular organelles are quite distinct in their glycerolipid composition (Jouhet etal., 2007). This reflects not only the different lipid biosynthetic pathways which contribute to organelle biogenesis, but also the unique functions that are carried out by these subcellular compartments. This is clearly seen in the chloroplast thylakoids that provide a matrix for the photosynthetic machinery, where the non-phosphorous galactolipids mono- and di- galactosyldiacylglycerol (MGDG and DGDG, respectively) are the predominant glycerolipid species (Dormann and Benning, 2002). Under normal growth conditions in Arabidopsis, MGDG and DGDG are restricted to the chloroplast membranes, and are primarily synthesized by two UDP-galactose (UDP-Gal) dependent galactosyltransferases, MGDI and DGD1(Benning and Ohta, 2005). MGDI is present on the inner chloroplast envelope and converts diacyl glycerol (DAG) derived from both the plastidic and ER glycerolipid biosynthetic pathways to MGDG (Benning et al., 2006, Kobayashi et al., 2007, Shirnojima et al., 1997). MGDG is then trafficked to the thylakoid, or the outer chloroplast envelope, where it can be further galactosylated by DGDI (Dormann et al., 1999). The combined activities of the galactolipid biosynthetic and lipid trafficking pathways give rise, in part, to sub-organellar membranes with defined lipid compositions during normal growth. However, during environmental stresses, such as phosphate deprivation, plants adjust their lipid composition in order to survive. The Arabidopsis null-mutant, dgdl, shows a 90% decrease in DGDG levels and severely stunted growth (Dormann et al., 1995, Dormann eta]., 1999). However, dgd] plants grown under 34 phosphate-limiting conditions accumulate DGDG to ~60% of the levels found in wild type plants (Hartel et al., 2000)—indicating that a DGDl-independent DGDG biosynthetic route exists in plants. In this regard, a set of functional MGD] and DGD] paralogs, MGD2/3 and DGD2 respectively, have been characterized at the molecular level and were shown to be coordinately regulated at the transcript level, with strong induction under Pi-limiting growth conditions (Awai et al., 2001 , Kelly et al., 2003, Kelly and Dormann, 2002). MGD2/3 and DGD2 have been localized to the chloroplast outer envelope membrane, and have been proposed to contribute to the biosynthesis of DGDG destined for extraplastidic membranes (Andersson et al., 2005, Kelly et al., 2003). Another class of galactolipid biosynthetic enzyme has been found in plants—the galactolipid:galactolipid galactosyltransferases (GGGTs), which through processive activity can give rise to oligogalactolipids. While the gene(s) encoding GGGT(s) have not been identified, GGGT activity was observed in isolated chloroplasts of the Arabidopsis dgdl dgd2 double null-mutant (Kelly et al., 2003). Regardless, oligogalactolipids are undetectable in whole-leaf lipid extracts from wild-type Arabidopsis (Xu et al., 2003), and DGDG was not detected in dng dgd2 double null- mutant plants (Kelly et a1. , 2003), suggesting that GGGT(s) contribute minimally to overall galactolipid biosynthesis under normal growth conditions. While much progress has been made towards the identification and characterization of genes involved in galactolipid biosynthesis, the regulatory factors that govern these pathways in plants remain unknown. Toward this end, a suppressor screen in a dgd] EMS mutagenized population recovered a mutant, dgsl-l (dgdl suppressor ]- 1). The dgsl-I mutant shows constitutive activation of alternative DGDG biosynthesis, 35 with DGDG accumulating to ~60 % of the level found in wild-type leaves. The dgsl -1 mutation has been mapped to a nuclear gene encoding a ~69 kDa protein of unknown frmction with two predicted trans-membrane spanning domains (hereafier referred to as DGSI), where it causes a G-to-A mutation resulting in a substitution of Asp457 to Asn (D457N) in the deduced protein sequence. Initial studies by Changcheng Xu showed that the over-expressed DGSI-C-terminal-GF P fusion protein was associated with the E" mitochondria. Interestingly, over expression of the dgsl-I-GFP cDNA resulted in a severe dominant negative growth defect accompanied by early leaf senescence, while over-expression of the DGSI-GFP cDNA did not (Xu et al., 2008). These results suggested that DGSI may be present in a protein complex, as dominant negative phenotypes are often associated with the poisoning of a protein complex with a mutant protein (Herskowitz, 1987, Leavitt et al., 1999). When hydrogen peroxide (H202, a reactive oxygen species [ROS]) levels in leaves were determined, the dgsI -1 -GFP transgenic lines showed a two-to-three fold elevation over wild-type levels. Likewise, in homozygous dgsl-I plants, H202 levels were moderately elevated in leaves and roots. Subsequent experiments revealed that treatment of dgdl plants with H202 resulted in an increase in the levels of MGD2/3 transcripts as well as the accumulation of DGDG. These results provided a potential link between the elevated H202 in dgsl-I and the phenotypic suppression of dgdl that this allele causes. To further investigate the possible role of DGSI in regulating galactolipid biosynthesis, a comparative study of the dgsl-I allele with a newly identified null T-DNA allele, dgsI-Z was initiated. 36 Materials and Methods Plant materials and growth conditions. Wild-type Arabidopsis plants (Col-2), and the dgsI-I and dgdl dgsl -1 mutants are as previously described (Xu et al., 2003, Xu et al., 2008). The dgsl-Z T-DNA insertion line (Sessions et a]. , 2002), SAIL_391_F040, was obtained from the Arabidopsis Biological Resource Center at Ohio State University. This line was selfed and tested for homozygosity (lack of segregation of the T-DNA marker). Homozygous dgdl dgsI-Z double mutant plants were obtained from the F2 seeds of a cross between dgdl and dgsl-Z homozygous mutant plants. Genotyping for dgdl homozygosity was performed by PCR with a dCAPS marker (Xu et al., 2003). Homozygosity for dgsI-Z was confirmed by the absence of a PCR amplicon when using the forward and reverse primers 5’-GAGGTGAGTGAACTAGAACATTCCATGAG-3’ and 5'-ACTCAACATAAAGCTAGAAAGCCCATTG-3’, respectively, which span the T-DNA insertion, and the presence of a PCR amplicon when the above reverse primer is used with a primer complementary to the left border of the T-DNA insertion (Alonso et al., 2003). In general, surface sterilized seeds were grown on agar solidified MS medium or soil as previously described (Xu et al., 2003). Bioinformatic analysis of DGSI. Full-length, deduced amino acid sequences were aligned with ClustalX (version 1.81), and generation of the bootstrapped phylogentic tree was performed using the PHYLIP software package as previously described (Mamedov et al., 2005). For transmembrane domain prediction the TMHMM program was employed (Krogh et al., 2001). 37 Generation of the DGSI antibody and immunoblot analysis. A central portion of the DGSI cDNA (GenBank accession no. AK118777; bp 1029—1503) encoding the amino acids between the two predicted transmembrane-spanning domains was isolated by PCR using primers 5'-ACTGGATCCTCTATATGGCTACTA-3' and 5'- CATGGTACCTCATAATGCAGCCAGAAT-3'. The resulting fragment was digested with BamHI and KpnI and inserted into pQE30 (Qiagen). The expression strains were routinely cultured overnight at 37°C in LB ampicillin (100 pg ml'l), inoculated at 1/250 dilution in LB ampicillin (100 pg mf‘) and cultured at 37°C. The protein was induced with 0.4 mM isopropyl-B-D-thiogalactopyranoside at an optical density at 600 nm (ODGOO) of approximately 0.4. At 0D600 0.7—0.8, cultures were cooled on ice and cells were harvested by centrifugation at 5000 g. Cells from 250 ml of culture were resuspended in 1 ml of wash buffer (50 mM Tris pH 7.5, 600 mM NaCl, 20 mM irnidazole) and lysed by sonication. The protein was purified from cell Iysate using an Ni-NTA agarose column (Qiagen) according to the manufacturer's instructions. Fractions with the highest protein content were combined, and the protein concentration was determined by the Bradford method (Bradford, 1976) using BSA as a standard. Rabbit polyclonal antiserum against denatured protein in acrylamide was raised by Cocalico Biologicals (Reamstown, PA, USA). The antibody was affinity-purified as previously described (Ermolova et al., 2003) using an affinity matrix consisting of 75 ug of the antigenic recombinant DGSI polypeptide fragment bound to a PVDF membrane. 38 Isolation of mitochondria and chloroplasts. Mitochondria were isolated from leaf tissue of three-week-old, soil grown A rabidapsis plants and purified using Percoll gradients according to Keech et al. (2005), while crude cauliflower mitochondrial preparations were performed as previously described (Douce et al., 1972). Intact chloroplasts were isolated from Arabidopsis leaves as previously described (Bessoule et al., 1995). For SDS-PAGE/immunoblot analyses, denatured crude leaf extracts were prepared as previously described (Martinez-Garcia et al., 1999), while protein fractions from purified organelles were denatured in a standard SDS-PAGE buffer (Laemmli, 1970). Protein concentration was quantified using the RCDC protein assay kit from Biorad (Hercules, CA, USA) with BSA as a standard. For Blue Native PAGE, solubilized proteins from cauliflower or Arabidopsis mitochondria were prepared using Triton X-100, dodecylmaltopyranoside, or Tween-20 as described in (Eubel et al., 2003) at 0.2, 0.33, and 1.0 mg detergent per mg protein (all at 25 mg protein ml"). The supernatant fractions after a 16,000 x g centrifirgation were removed and mixed with Coomassie Brilliant Blue R-250, and 50 ill from each sample was separated by Blue Native PAGE (Eubel et al., 2003, Schagger et al., 1994). lrnmunoblotting was performed on PVDF membranes using the CPS-1 Chemiluminescent peroxidase substrate detection system from Sigma (St. Louis, MO, USA). Monoclonal antibodies against a conserved epitope in plant alternative oxidases (AOX mAb) (Finnegan et al., 1999) were obtained from stock at the Department of Energy Plant Research Laboratory, Michigan State University, left by the late Lee McIntosh. Monoclonal antibodies against the outer mitochondrial membrane VDAC protein were provided by Tom Elthon (University of Nebraska-Lincoln, USA). Polyclonal antibodies against the pea chloroplast outer 39 envelope membrane protein TOC 75 were kindly provided by John Froehlich (Michigan State University, USA) (Tranel et al., 1995). RNA Analysis. Total RNA from liquid N2-frozen leaf tissue of two-week-old seedlings was isolated with the QIAGEN RNeasy Plant Mini-kit (QIAGEN, Germantown, MD) according to the manufacturer’s instructions and separated by electr0phoresis through a formaldehyde-denaturing agarose gel. RNAs were then transferred to a nylon filter which was processed for RNA blotting using ULTRAhyb hybridization buffer (Ambion, Austin, TX, USA). The probe used to specifically detect AOX 1 a mRNA was generated by PCR with the following primers: forward, 5’-TTTGTTCTTCGACGGTCCGTAC-3’; reverse, S’CGCGATTCCTFTATCTCCCT‘TG-T, and labeled with ”P using the Megaprime DNA Labeling system (GE Healthcare, USA). Total RNAs were stained with methylene blue as previously described (Wilkinson et al. , 1991). Lipid Analysis. Lipids were extracted from leaves (Dormann et al., 1995) on an equal fresh weight basis and separated by thin-layer chromatography on activated ammonium sulfate-irnpregnated silica gel TLC plates (Si250, Mallinckrodt Baker, NJ, USA) with a solvent system of acetone/toluene/water (91 :30:7.5, by vol.). Gas chromatography based quantification of fatty acid methyl esters (FAMEs) was performed as previously described (Xu et al., 2005). MGDG and DGDG bands were scraped into individual vials, while the remaining polar lipids were collectively scraped into a single vial for FAME derivatization (designated as the remainder of polar lipids). Lipids were extracted fi’om mitochondria as described above, but loaded on TLC plates on an equal total 40 mitochondrial protein basis. For visualization of total lipids separated by TLC, dried plates were placed in a chamber containing 12 vapors; for visualization of glycoglycerolipids, plates were stained with a-naphthol (Benning et al., 1995). Results Introduction of dgsI-2 does not suppress the dgdl growth phenotype. To further characterize the function of DGS I , homozygous plants were generated for a potential null T-DNA allele (SAIL_391_FO40 (Sessions et al., 2002)) inserted into the third exon of DGSI (AT5G12290) and confirmed by PCR (Figure 2.1A). This T-DNA allele, designated dgsl-2, results in an insertion of the left border of the T-DNA plasmid at a codon corresponding to the 159th amino acid—which is N-temrinal to the D457N mutation caused by dgsl -1 (Figure 2.1B). Analysis of 30-d-old plants revealed no obvious growth defects in the dgsl-l or -2 mutants compared to wild-type, where as these alleles had distinct phenotypes when observed as double mutants in a dgdl background: dgsl -1 suppressed the growth defect of dgdl, whereas dgsI-2 had no effect (Figure 2.1C). This result is consistent with the hypothesis that dgsl -1 is a gain-of function allele, whereas dgsI-Z is a loss-of-firnction allele. Moreover, the lack of a clear phenotype for dgsI-2 in a wild-type background indicates that DGSI is not essential for normal growth and development. DGSI is a mitochondrial membrane protein. DGSI encodes a 602 amino acid, ~69 kDa protein of unknown function. DGSI is predicted to contain two transmembrane- spanning domains when analyzed with the TMHMM 2.0 program (Keech et al., 2005) (Figure 2. I B), and a BLAST search (Altschul et al., 1997) revealed several DGSI 41 A RP+LP LB1+LP B M 1 2 1 2 dgs1-2 (T-DNA) 3%217111 V59 Lo TM1 TM2 Figure 2.1. Comparison of the dgsI-Z T-DNA null allele and the dgsI-I point mutant gain of function allele in wild type and dgdl backgrounds. (A) Genotyping of wild- type (1) and dgsI-Z homozygous (2) plants using the PCR primers RP, LP and LB] described under MATERIALS AND METHODS. (B) Diagram showing the relative positions of the dgsl -1 and dgsl-Z mutations in the DGSI protein. The relative positions of two predicted transmembrane spanning domains are also indicated (TMI and TM2). (C) Growth phenotypes of 30-day-old plants sown directly on soil. The respective genotypes are indicated along the bottom. 42 Athaliana pIHrrFs E. DQILRANEI AI A.oryzae PPVGT R DI ID-LLKSQEL Gr e.glabramPIKNIVT S IDKMLKSQQL GI c. albicansPIKYLIT S IDKLLKSQQL AM c.grobosumPIVGMK D ID-LLKSQEL or admoideum PLVGAVS D IDSLLKSQEL GF QNL K a DQILKANEI AI E‘ftfiifl’ Qurllr D IDKLLQANEL QL Ow," Rim. T D IDKLLQSQQL GI ' . KGLSS K DQILRANEI AI O'ta‘mPLTt-JNL D DQLNRANRL SL P-"O'dumprrx'car N IDSILKSQEL GE‘ P-"lChOCfifPapIRrav N IDRLLKSQEL AT S-cerewaaeprasar N DEVLEGNDL KI s.pombe PVKSLI s IDQILKSQEL GI Y. llpolytica IKNQ G SLLIQLQK IDKMLKSQQL Gv C. albicans E. gossypii - - S. cen'visae 7. cruzi Y. Irpolyttca + + C. glabrata + 0 S. pombe 9 a + A. oryzae a. discoidium 2 p nommm + N. crassa + C. globosum P. tn'chocarpa + + A. thaliana 0.1 O. sativa O. taun' Figure 2.2. Sequence analysis of DGSI. (A) Partial sequence alignment of DGSI orthologs showing the region of highest identity. Conserved residues are shaded in black and the asterisk indicated the Asp residue mutated to Asn in dgsl (D457N). (B) Unrooted tree showing the relatedness of predicted DGSI orthologs in plants and fungi. Boot strapping values of >950 are marked by +. Those between 500 and 900 are marked with a solid circle. 43 orthologs in plant, algal, and fungal species—as well as other organisms such as Dictyostelium discaideum and Ttypanasoma cruzi—but not in animals or prokaryotes (Figure 2.2). Intriguingly, the Asp457 residue that is mutated in the dgsl-1 allele (D457N) is entirely conserved among the DGSI orthologs in a region of high local similarity (Figure 2.2A). The DGSI protein is predicted to be targeted to the mitochondrion or chloroplast according to the Arabidopsis subcellular database (SUBA) (Heazlewood et al., 2007 ), and was identified in the Arabidopsis mitochondrial proteome (Heazlewood et al., 2004). The fluorescence of a C-terrninally GFP-tagged DGSI fusion protein was associated with organelles consistent with the appearance of mitochondria, but not chloroplasts (Figure 2.3A). The localization of the DGSI-GFP fusion protein was further confirmed by overlap of the corresponding GFP fluorescence with Mitotracker Red fluorescence in root hairs of the DGSI-GFP transgenic plants (Figure 2.3B). Irnunodetection of DGSI with a specific polyclonal antibody generated in this study revealed that the protein could only be detected in isolated Arabidopsis mitochondria, but not crude leaf, or isolated intact chloroplast preparations (Figure 2.3C). The lack of a signal in the mitochondrial fraction when using an antibody against the outer chloroplast envelope protein TOC75 ruled out chloroplast envelope contamination. Together, these results suggest an exclusive localization of DGSI to mitochondria. Recent proteomic studies on mitochondrial membrane fractions from yeast and Neurospora identified homologs of DGSI in outer mitochondrial membrane preparations (Schmitt et al., 2006, Zahedi et al., 2006). To determine the sub-mitochondrial membrane association of DGS 1 , mitochondria from cauliflower (Brassica oleracea) florets were g, - kDa 180- 115- g .. 82 , 2 § .. 64- 24‘2” ‘: DGSt-GFP Mitotracker Merged 3, a Red 49- El. ...... CBB w- . "U“ C K 37- . .. (b \ .\° 0 ‘0 \ ,. V O ‘1‘ ‘ fl $\°\\°\ 254:-afi.‘ v‘ DGS] 19' ‘ 15- ! ._. —- - C" AOX — TOC7S Figure 2.3. Localization of DGSI in mitochondria. (A) Confocal microscopy image of Arabidopsis wild type stably expressing a DGSI-GFP cDNA. The images for chlorophyll fluorescence (red) indicative for chloroplasts, and for GFP indicative for the presence of the DGS 1 -GFP fusion protein (green), were overlaid. The scale bar equals 8 pm. (B) Confocal microscopy images of a single root hair of Arabidopsis wild type stably expressing a DGS] -GFP cDNA; the GFP and Mitotracker Red fluorescence images are shown separately, as well as the merged image as indicated. The scale bars equal 10 pm. (C) Irnmunodetection of DGSI in isolated Arabidopsis mitochondria. Proteins extracted from wild-type Arabidopsis leaf (Wt leaf), intact chloroplasts (Int Chl) and intact mitochondria (Int Mito) were separated by SDS—PAGE (50, 80 and 15 pg, respectively), and immunoblotted with DGSI , AOX (a mitochondrial inner membrane protein) and TOC 75 (a chloroplast outer envelope protein) antibodies as indicated. (D) Association of DGSI with sub-mitochondrial fractions from cauliflower. Immunoblots with lanes containing total protein (Tot Prot), intact mitochondria (Int Mito), outer mitochondrial membrane (OMM) and inner mitochondrial membrane (IMM) are shown. Three identical blots were probed with antibodies against DGS 1 , VDAC (an outer membrane protein) and AOX (an inner membrane protein). Equal amounts of protein were loaded as indicated by staining a gel run in parallel with Coomassie Brilliant Blue (CBB), where apparent molecular weights of an included protein marker set are indicated (kDa). 45 isolated and inner and outer membrane fractions were prepared. The Arabidopsis DGS 1 - specific antibody detected a protein of the correct size (Figure 2.3D) as well as a lower molecular weight protein that was not further investigated. As a marker for the outer membrane, the voltage-dependent anion channel (VDAC, porin) was used; alternative oxidase (AOX) was included as an inner membrane marker. DGSI co-purified with the outer mitochondrial membrane marker VDAC (Figure 2.3D), and was clearly not enriched in the inner mitochondrial membrane fraction as was observed for AOX. These results are consistent with the presence of DGSI orthologs that have been localized to the outer mitochondrial membranes of fungi, but co-localization to the inner membrane cannot entirely be ruled out. DGSI is present in a high molecular weight complex. One of the simplest mechanisms—and a common underlying factor—that gives rise to dominant negative phenotypes is the. poisoning of a protein complex by a mutated component protein (V eitia, 2007). To test whether complex poisoning was a possible mechanism that underlies the dgsl-1 phenotype, we explored whether the DGSI protein is present in a higher M, complex. Initially, mitochondria isolated from cauliflower florets were used as a facile source that provides high quantity without contamination of chloroplast membranes. Following solubilization of mitochondrial proteins using increasing amounts of Triton X-100 and separation of complexes by blue-native PAGE, a high Mr complex migrating between the 440 and 669 kD markers (approximately 500 kD) containing a peptide reacting with anti-Arabidopsis DGSI antibody serum is present (Figure 2.4A). This complex is stable up to 1 mg Triton-X100 per mg protein, the highest concentration tested. As a control, complex IV was monitored using a monoclonal antibody that 46 Figure 2.4. Blue Native PAGE analysis of the DGS] complex. (A) Cauliflower mitochondrial proteins solubilized with varying amounts of Triton X-100 (0.2, 0.33, and 1 mg Triton X-100 per mg mitochondrial protein in lanes 1, 2 and 3, respectively) and separated by blue native PAGE. The Coonrassie Brilliant Blue stained proteins (CBB) and corresponding immunoblots with the DGSI antibody (DGSI) or the COXII antibody (COXII) are shown. The relative positions of protein standards are indicated (thyroglobulin at 669 kD, ferritin at 440 kD, and catalase at 200 kD). (B) Blue Native PAGE of cauliflower mitochondrial proteins solublized with different detergents Triton X-100 (lane 1), Dodecylmaltopyranoside (lane 2), or Tween 20 (lane 3), followed by immunodetection of DGSI. (C) Blue Native PAGE separation of Triton X-100 solubilized crude cauliflower mitochondria (lane 1), and from isolated Arabidopsis mitochondria from wild-type (lane 2) and homozygous dgsl-1 plants (lane 3). 47 recognizes subunit II (COXII), and was present in a high M, complex (approximately 300 kD) presumably representing complex Vla of the respiratory electron transport chain as previously described (Eubel et al., 2003). Contrary to the DGSI complex, this complex was not stable at 1 mg Triton-X100 per mg protein, where only the previously described (Eubel et al., 2003) complex M was detected (Figure 2.4A). The DGSI immunogenic complex was not affected by using other detergents, dodecylmaltoside or Tween 20, for solubilization (Figure 2.48). Isolated leaf mitochondria from Arabidopsis also showed the presence of a DGS l-immunogenic complex similar in apparent M, to the cauliflower DGSI complex (Figure 2.4C). The Arabidopsis DGSI complex was indistinguishable in wild-type and dgsl-1 mitochondria, indicating that the DGS 1 04573; protein produced in dgsl -1is not defective in complex formation. These data clearly indicate that DGSI is present in a multimeric complex, but do not provide any evidence as to whether DGSI is interacting with other proteins (heteromultimer), or forms a homomultimeric quaternary structure. Phosphate deprivation and dgsl-1 allelism have additive effects on lipid metabolism. Phosphate deprivation has been reported to lead to accumulation of DGDG in mitochondria of plant cells (Jouhet et al., 2004). As DGSI is a mitochondria-localized protein (Figure 2.3), potential changes in mitochondrial lipid composition in dgsl-l plants were investigated. As shown in Figure 2.5 no differences in lipid composition were observed between mitochondria isolated from wild-type, dgdl , or dgdl dgs1-1 mutant leaves from plants grown under normal conditions. Irnportantly, no DGDG was found in the dgdl dgsl-1 mitochondria, indicating that the DGDG accumulating in these mutants is associated with non-mitochondrial membranes. As such, these results suggest that the 48 A 3‘ A ,t .3 39 K N N N $9 efil a? ~§K 89 $9 r— whole leaf --1 ,— mitochondria—I MGDG ' PG/DPG ' " DGDG — SQDG Pl/PS PE PC MGDG O - .- DGDG ’ SQDG TGDG ........................ Figure 2.5. Mitochondria of dgdl dgsl-I homozygous mutant plants do not accumulate galactoglycerolipids. Lipid extracts were prepared from leaves (on an equal fresh weight basis) and isolated mitochondria (on an equal mg protein basis) separated by TLC and stained with iodine (A), and after de-staining, with a-naphthol (B). Lipids: DGDG, digalactosyldiacyl glycerol; DPG, diphosphatidylglycerol (cardiolipin); MGDG, monogalactosyldiacyl glycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; SQDG, sulfoquinovosyldiacylglycerol; TGDG; trigalactosyldiacyl glycerol. 49 processes of DGDG accumulation due the presence of the dgsl-I allele and during phosphate-limiting growth conditions may be distinct. To test whether DGSI is required for DGDI -independent DGDG biosynthesis following phosphate deprivation, galactolipid content was determined in the dgdl single mutant and the dgdl dgsl-1 and dgdl dgsl-2 homozygous double mutants under normal growth conditions (Figure 2.6A) and under conditions of phosphate deprivation (Figure 2.6B). The dgdl dgsl-2 double mutants were indistinguishable from the dgdl mutant with regard to DGDG accumulation under phosphate deprivation, suggesting that DGSI is not required for phosphate-dependent activation of the DGDI-independent pathway. The accumulation of DGDG in the dgdl dgsl-1 homozygous double mutant under phosphate deprivation was higher than in dgdl, indicating that dgsl-I is additive. This result provides further evidence that phosphate deprivation and introduction of the dgsl-I mutation act independently and represent distinct mechanisms of activating the DGD l - independent pathway(s) of galactolipid biosynthesis. In addition to the paralog of DGD], DGD2, which is known to be induced following phosphate deprivation (Kelly and Dormann, 2002), a distinct processive galactolipid:galactolipid galactosyltransferase (GGGT) discovered in preparations of plastids (van Besouw and Wintermans, 1978) but not yet defined at the molecular level might be activated in dgsl-1 as well. This appears to be the case, as analysis of leaf lipid crude extracts by TLC revealed the accumulation of the oligogalactolipid trigalactosyldiacyl glycerol (TGDG) in the dgdl dgsl-l double mutant following phosphate deprivation (Figure 2.6 C). TGDG is a product of the above-mentioned GGGT 50 Figure 2.6. Galactoglycerolipid content in the dgdl single homozygous mutant and the dgdl dgsl-1 and dng dgsl-2 double homozygous mutants under normal and phosphate-deprived growth conditions. Monogalactosyldiacylglycerol (black bars) and digalactosyldiacylglycerol (grey bars) relative amounts present in leaves of 10—day-old plants grown on MS agar plates (A), and (B) 10 d old plants transferred to MS agar plates lacking phosphate for an additional 10 days. Relative amounts are expressed on a % mol basis of polar glycerolipid and the average and standard deviation of three independent measurements is shown. (C) A TLC plate stained with u-naphthol to reveal the separated galactoglycerolipids. Plants were grown in the presence (+) or absence of phosphate (-) as described for (A) and (B). DGDG, digalactosyldiacylglycerol; MGDG, monogalactosyldiacylglycerol; TGDG, trigalactosyldiacylglycerol. 51 A dgd1 dgs 1-2 dgs1-1 dgd1 dgd1 dgd1 dgs1-2 dgs1-1 dgd1 dgd1 000000 00000 54321 54321 0 $225 8:... :28 gag 8.2. 3.8 B C G D G D McDG" 52 and is also found in the tgd mutants defective in endoplasmic reticulum-to-plastid lipid trafficking (Xu et al., 2003). TGDG also accumulates in ozone-treated spinach (Spinacea aleracea) leaves (Sakaki et al., 1990). This may be related to the ROS accumulation in dgsl—I mutant plants (Xu et al., 2008), which would also be produced following ozone treatment. The fatty acid profile of MGDG and DGDG in the three different mutant lines showed no difference between dgd1 and the dgdl dgsl-2 homozygous double mutant line under phosphate replete and depleted growth conditions (Figure 2.7). The dgd1 dgsl-1 homozygous double mutant showed slight differences from dgd1, with a slight increase in 16:3 (carbons: position of double bonds) and decrease in 18:3 fatty acids in MGDG and lowered relative amounts of 16:0 and increased 18:3 fatty acids in DGDG independent of the growth conditions. Alternative oxidase protein levels are reduced in the dgsl-1 mutant allele. In order to determine if defects in mitochondrial protein composition are present in the dgsl-I mutant plants, the protein levels of DGSI , alternative oxidase (AOX), and COX II subunit of complex IV in isolated mitochondria were investigated. Mitochondria were prepared from leaves of the wild-type, the dgsl-2, dgd1, dgsl-1, and the dgd1 dgsl-1 homozygous single and double mutants, respectively, and protein levels were compared using immunoblotting. As shown in Figure 2.8, the dgsl-2 T-DNA allele results in the ablation of the full length DGSI protein, confirming that this is a null or loss-of-function allele as indicated above. Moreover, the presence of the dgsl-I allele does not lead to a 53 Figure 2.7. Galactolipid Fatty acid profiles. Fatty acid profiles of mono- (MGDG; A, B) and digalactosyldiacylglycerol (DGDG; C, D) in the dgd1 single homozygous mutant and the dgd1 dgsl-1 and dgd1 dgsl -2 double homozygous mutants under normal (A, C) and phosphate-linrited (B, D) growth conditions. The data are presented on a % mol basis and represent the average and standard deviation from three independent experiments. Fatty acids are designated with carbon numbers : number of double bonds. 54 C70 Fatty Acid (mol %) o a a a a a a Fatty Acid (mol %) U A N or b 0'! O) N O O O O O O O O Idgdt , I dgd1 ndgdt dgs1-1 f ‘ 16:0 16:1 ' Idgdt u dgd1 dgs1-2 adgd1 dgst-t dgs1-2 MGDG (+Pi) 16:2 16:3118207182‘1'182 18:3 MGDG (-Pi) 18:3 16:0 16:1 16:2 16:3 18:0 18:1 18:2 ‘ Idgdt ‘ Edgd1 dgs1-2 _ Idgdt dgst-t DGDG (+Pr) 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 ' Idgdt adgdt dgs1-2 . ldgdt dgs1-1 DGDG (-PI) A a I A 7 I A A 16:3 18:0 18:1 18:2 18:3 55 *“um- __ one “m “'"‘ ""5 ""‘W Stain . I. ~ C .04.». _ H pow m “_ Figure 2.8. Alternative oxidase is reduced in dgsl-1 mutant mitochondria. Immunoblots of proteins from isolated mitochondria prepared from leaves of wild type (WT), dgsl-1, dgsl-2 and dgd1 single homozygous mutants, and the dgd1 dgsl-1 double homozygous mutant are shown. The blots were probed with affinity purified DGSI antibody (DGSI), an alternative oxidase antibody (AOX), or a cytochrome c oxidase subunit II antibody (COX 11). Equal amounts of protein were loaded as shown in the Coomassie Brilliant Blue (CBB) stained gel at the bottom. 56 decrease in DGS I-immunogenic protein; if anything the protein level might actually be slightly increased in dgsl-1. An AOX monoclonal antibody that detects all major forms of this protein (Finnegan et al., 1999) was initially included as a potential loading control for the experiment. However, it became apparent that AOX levels are strongly reduced in the dgsl-l gain-of-function mutant but unaffected in dgsl-2 (Figure 2.8). The observed loss of the AOX protein was not due to a general deterioration of mitochondrial complexes as COX II was unaffected in any of the mutant backgrounds (Figure 2.8). Moreover, a corresponding Coomassie Brilliant Blue-stained gel (Figure 2.8, bottom section) did not show any major differences in the mutant lines, suggesting that there is no large-scale aberration in mitochondrial protein patterns in the mutant backgrounds analyzed. Potential defects in the expression of the AOX1a gene of Arabidopsis were examined. AOXIa is the dominantly expressed AOX encoding gene in Arabidopsis seedling tissue. To be able to readily detect AOXla transcript levels in seedling shoot tissue samples, we treated the seedlings with Antimycin A, which specifically inhibits respiratory electron transport at complex III and leads to a transient induction of A 0X 1a expression (Zarkovic et al. , 2005). As shown in Figure 2.9, the presence of neither of the two dgsl mutant alleles affects induction of A 0X1 a mRNA expression. As such, the reduction of AOX protein levels observed in isolated mitochondria from dgsl-1 mutant plants must be due to a posttranscriptional mechanism. Moreover, the wild-type DGSI protein is not involved in regulating AOX protein or transcript levels, at least under the conditions tested, because the dgsl-2 loss-of-function allele had no effect. The defect in 57 WT dgs1-2 0246810122402468101224 Btu-"911.0 nuns-cu on1a lam—‘I— an... .— I an..." — - mun—Ar- mRNA —--— . . ,u-u- .- - .. .u . . it...biOv ,. ore-O...’""—.‘“ l lbaeaclitocoouuoo.4eoilbaour D’.‘." :f' " -t.:.-.s,. Lvr...a.a~ ... _.‘._.,.-.-. x ‘_=!=-A‘Ilu" " F~.\er-.I-~ ‘~ -.4 . 3""! l-a . a." — I ' am .n' '0'" "' 1" W '-' Ill WT dgs1-1 0246810122402468101224 unit-Diva Hil-IfiiiHonta m-r-a can“ a... - .H. Witt-n .ygeauuuuHNu—uflnuu ......auaea-«IOIanIQOOueOduso rRNA ‘ -' Iaaoursla..a.... -: rarer- Figure 2.9. Analysis of AOXIa transcript levels in seedlings treated with Antimycin A. Two week-old seedlings of the wild type (WT) and the dgsl-l and dgsl-2 homozygous mutants were sprayed with 25 pM Antimycin A and samples of aerial tissue were taken at the indicated times (0-24 h). Total RNA (2.5 pg) from these samples were isolated, processed for RNA blotting and specifically probed with for A OXla mRNA Methylene Blue stained total RNA (rRNA) is shown as a loading control. 58 AOX protein in dgs] -1 is thus an additional phenotype specific to this gain-of-function allele as was observed for the above-mentioned lipid phenotypes. The dgsl-I mutant shows phenotypes of an AOXla-deficient mutant. Arabidopsis Mutants deficient in AOX generally have subtle biochemical and morphological phenotypes under normal that become more apparent only when exposed to multiple environmental stress conditions (Giraud et al., 2008, Strodtkotter et al., 2009, Umbach et al., 2005). One easily observable phenotype is reduced root growth of AOXla-deficient lines (Giraud et al., 2008). Comparing root length of seedlings between the different Arabidopsis lines used in this study, only lines carrying the dgsl-1 allele have shorter ' roots when compared to the respective wild-type or dgdl mutant backgrounds (Figure 2.10). Thus, the dgsl-I mutant plants, which have reduced protein AOX protein levels phenocopy the reduced root growth observed in AOXla-deficient T-DNA lines. While this suggests that some of the phenotypes observed in dgsl-I might arise from the AOX protein defect, there are differences in phenotypes observed between these two mutant classes: dgsl-l plants accumulate hydrogen peroxide (Xu et al., 2008), which was reported not to be the case for the AOXla-deficient lines (Giraud et al., 2008). 59 Root length (mm) _a N (A) «b 0'1 0’ O O O O O O O a WT dgs1-2 dgs1-1 dgd1 dgd1; dgd1: dgs1-2 dgst-t Figure 2.10. The dgsl-I but not the dgsl-2 mutant displays a root growth phenotype. Seedlings were grown for 10 d on vertically placed MS agar plates supplemented with l % sucrose, and root length was measured. The data represent the average and standard deviation from 5 individual plants. 60 Discussion This study was initiated to determine whether the DGSI protein plays a direct role in the regulation of DGD] -independent DGDG biosynthesis. Initial studies focused only on the gain-of-function dgsl-1 allele (Xu et al., 2008) which causes severe growth defects upon stable over-expression. The DGSI protein was implicated as a potential factor involved in the regulation of alternative DGDG biosynthesis due to the partial suppression of DGDG deficiency observed in the dgdl dgsl-1 double mutant, which is similar to the effect of phosphate-deprivation induced DGDG accumulation (Hartel et al., 2000, Kelly and Dormann, 2002). Given the increased H202 and reduction in AOX protein levels in dgsl-1 (Figure 2.8), and that the DGDI -indepdendent pathway is induced by exogenous H202 application (Xu et al., 2008), one could conceive a simple hypothesis for the possible function of DGSI in phosphate-mediated regulation of galactoglycerolipid biosynthesis. Phosphate deficiency is known to lead to a slowing of ATP synthesis in mitochondria (Theodorou and Plaxton, 1993), causing a backup of the electron transport chain as the two processes are tightly coupled. The resulting over-reduction of the respiratory electron transport chain leads to ROS formation that can be compensated for by AOX activity, which can directly transport electrons from ubiquinone to oxygen. If DGSI regulated AOX activity or abundance in response to phosphate deprivation, a mechanism by which ROS could serve as a signal for the activation of the DGDl- independent pathway can be envisioned. However, two observations from this study disagree with this hypothesis: (i)dgs1-2 mutants completely lack DGSI but are indistinguishable from wild-type in their response to phosphate deprivation with regard to galactolipid metabolism at the level tested, and (ii) DGDG accumulation due to the 61 presence of the dgsl -1 allele and phosphate deprivation are additive and therefore involve two independent mechanisms. Based on these observations it can be concluded that DGSI is not directly involved in the low phosphate induction of DGDG biosynthesis by the DGDl-indepdendent pathway. Regardless, this work has revealed a clearer understanding of how the dgsl-l allele may cause the activation of DGDI-independent DGDG accumulation. As previously observed, this pathway is induced by ROS, which is present in elevated levels in dgsl -1plants (Xu et al., 2008). In both dgsl-1 and H202-treated wild-type plants, transcripts are elevated for the MGDG synthase encoding paralogs of MGD] : M602 and MGD3 (Xu et a]. , 2008). The accumulation of TGDG in phosphate-deprived dgd1 dgs]- ! double mutants indicates that GGGT, which remains to be identified at the molecular/genetic level, is also contributing to DGDG biosynthesis in dgsl-1. The role of GGGT in lipid biosynthesis is poorly understood, but is known not to contribute significantly to DGDG biosynthesis under normal conditions (Kelly et al., 2003). GGGT is activated in the tgd lipid trafficking mutants in Arabidopsis (Xu et al., 2003, Xu et al., 2005) and in ozone-treated spinach (Sakaki et al., 1990). The conditional activation of GGGT in dgsl-1 further demonstrates the importance of identifying the GGGT encoding gene(s), before a more complete picture of galactolipid biosynthesis can emerge. The reduction of AOX protein in dgsl-1 mutants seems a plausible cause of the observed accumulation of hydrogen peroxide in these plants. Indeed, a role for AOX in the alleviation of mitochondrial ROS has been well documented (Fiorani et al., 2005, Maxwell et al., 1999, Umbach et al., 2005). This conclusion appears to be in conflict with a recent report that AOX l a-deficient mutants do not accumulate hydrogen peroxide 62 (Giraud et al., 2008). However, a more recent report suggests that one of the other (five in total) AOX encoding paralogs in Arabidopsis, AOX 1d, is induced in the AOXla mutant background and may partially compensate for AOXla deficiency (Strodtkotter et al., 2009). Thus, at this time the question of whether the reduction of AOX protein observed in dgsl-1 is the underlying cause of hydrogen peroxide accumulation in this mutant line remains. How the dgsl-l mutant protein affects AOX protein levels also remains unresolved, but based on our AOX l a transcript analysis the effect must be posttranscriptional. Because DGSI is present in the outer membrane (Figure 2.3), while AOX is associated with the inner membrane, direct interactions between the two seem unlikely. It is also not likely that the DGSI wild-type protein is required for the processing or import of AOX protein into the mitochondria, because AOX protein levels are not affected in dgsl-2 plants. Moreover, the presence of DGSI orthologs in organisms that lack AOX (e.g., Saccharamyces cerevisiae) also suggest that DGSI is not necessarily involved in the regulation of AOX. Thus, the effect of the dgsl-I allele on AOX protein levels may not be directly related to the function of the DGSI wild-type protein. This work has revealed that the DGSI protein is not directly involved in the regulation of DGDl-independent galactolipid biosynthesis during phosphate deprivation, as originally hypothesized. The reduction in AOX protein also seems to be an indirect effect of the dgsl -l mutation, and if not directly involved in either of these processes, what is the molecular function of DGS l ? This question has remained difficult to answer because analysis of dgsl-2 has lead to no discemable phenotypes under the stress conditions tested. The biological function at the organismal level of many Arabidopsis 63 proteins including well-studied examples, such as AOX, is not clear as well. Many plant proteins are likely to be of conditional importance, and help sessile plants to cope with a number of environmental stresses. Indeed, the conditions under which the loss of the DGSI protein becomes critical have not yet been identified. 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(2006) Proteonric analysis of the yeast mitochondrial outer membrane reveals accumulation of a subclass of preproteins. Mal. Biol. Cell 17, 1436-1450. Zarkovic, J., Anderson, S. L., and Rhoads, D. M. (2005) A reporter gene system used to study developmental expression of alternative oxidase and isolate mitochondrial retrograde regulation mutants in Arabidopsis. Plant Mol. Biol. 57, 871-888. 68 Chapter 3 Identification of the gene encoding galactolipid:galactolipid galactosyltransferase and its role in freezing tolerance 2 The data represented in figures 3.2 and 3.8 are derived from experiments conducted by Bagyalakshmi Muthan. 69 Abstract UDP-galactose independent biosynthesis of di-, tri-, and tetra-galactoglycerolipids (DGDG, TGDG, and TeDG, respectively) has long been known to occur in plants. This distinct pathway of galactolipid biosynthesis is carried out on the chloroplast outer envelope by galactolipid:galactolipid galactosyltransferase (GGGT). GGGT transfers a galactosyl moiety from one MGDG molecule to another, forming DGDG and diacylglycerol, and through processive activity produces oligogalactolipids (TGDG and TeDG). Prior to this study, the gene(s) encoding GGGT were not known, and the role of GGGT in lipid metabolism remained enigmatic. Here, the GGGT encoding gene is identified as SENSITIVE T0 FREEZING 2 (SFR2), a gene of unknown function, which was previously identified when sfiz mutants showed freezing sensitivity. In viva and in vitro studies indicate the enzyme is activated by stresses including freezing, dehydration, and hyperosrnotic treatments. T-DNA mutants in SFR2 cannot produce oligogalactolipids and exhibit sensitivity to freezing, suggesting a role for GGGT in stabilizing the outer envelope during this stress. The SFR2 protein shows GGGT activity when expressed in yeast. Together these results indicate that GGGT is activated under sub-zero temperatures and remodels galactolipids in the chloroplast outer envelope—possibly by direct mechanosensation of physical changes that occur in the membrane during freezing. 70 Introduction The biosynthesis of glycerolipids has been intensely studied, not only because these molecules are important building blocks for biomembranes that provide the boundaries of the cell and organelles, but also because membranes are the site of signal and energy transduction processes that often require distinct lipid compositions for proper function. Molecular genetics has been a powerful tool in elucidating the factors required for glycerolipid biosynthesis in plants, where classical biochemical approaches to purify what are often integral membrane enzymes has been of more limited success. One example of this is the identification of the enzyme and encoding gene, DGD], responsible for the bulk of DGDG biosynthesis in Arabidopsis. The dgd1 null mutant exhibits a severe growth defect, and has a ~90 % decrease in DGDG levels in comparison to wild-type (Dormann et al., 1995, Dorrnann et al., 1999). Analysis of DGDl revealed it to be a UDP-galactose (UDP-Gal) dependent enzyme which acts on monogalactosyldiacyl glycerol (MGDG) as a lipid substrate. These findings were in seeming conflict with earlier biochemical studies, which suggested that further galactosylation of MGDG to form DGDG was carried out in a UDP-Gal independent manner by the unidentified enzyme galactolipid:galactolipid galactosyltransferase (GGGT) (Heemskerk et al., 1990, van Besouw and Wintermans, I978). GGGT catalyzes the transfer of a galactosyl moity from one MGDG molecule to another, forming diacylglycerol (DAG) and DGDG. Through processive activity, DGDG can be further galactosylated, producing oligogalactolipids (TGDG and TeDG) (Benning and Ohta, 2005, Heemskerk et al., 1983, van Besouw and Wintermans, 1978). 71 More recent studies in Arabidopsis revealed a DGD] paralog, DGDZ, which comprised an alternative pathway for DGDG biosynthesis that is activated at the transcript level upon phosphate deprivation. Intriguingly, the dgdl dgd2 double mutants had no detectable levels of DGDG, but chloroplasts isolated from these mutants were not defective in radiometrically measured GGGT activity (Kelly et al., 2003). These results clearly indicated that a distinct gene is encoding GGGT, and that GGGT does not play a significant role in DGDG biosynthesis under normal growth conditions. The role GGGT plays in galactolipid metabolism has remained unresolved, but other studies have indicated that GGGT is activated in response to ozone treatment in spinach leaves (Sakaki et al., 199%, Sakaki et al., 1990c), and also in the lipid trafficking defective tgd mutants in Arabidopsis (Awai et al., 2006, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008), indicating that GGGT may be involved in stress responses. This study was initiated to identify the GGGT encoding gene using a facile mutant screen in Arabidopsis, where infiltration of plant leaves with MgCl2 results in the accumulation of oligogalactolipids. Initial studies focused on finding mutants defective in oligogalactolipid accumulation in an ethane methyl sulfonate (EMS) mutagenized population of Arabidopsis. However, it was an alternative approach screening T-DNA lines of candidate genes—selected on the basis of the localization of the encoded protein and its potential to catalyze the GGGT reaction (glycosyltransferases and glycosyl hydrolases)——which lead to a candidate GGGT encoding gene. 72 Materials and methods Plant growth conditions. Plants were grown on soil or agar solidified Murashige and Skoog (MS) medium Murashige and Skoog, 1962) containing 1 % sucrose as previously described (Xu et al., 2005) on a 16 h light] 8 h dark photoperiod. Cold acclimation was achieved by placing the plants in a cold room at 7 °C at a photosynthetic photon flux density of 75 pmol rn'2 sec’2 with a 12 h 1i ght/ 12 h dark photoperiod. Freezing conditions were imposed by incubating plants at -2 °C (24 and 12 h total for MS-agar and soil grown plants, respectively) with ice nucleation after 3 h (Li et al., 2008). This was followed by lowering the temperature at 1 °C h’1 to -6 °C and incubation for 24 h. For post freezing recovery, plants were returned to the above-mentioned cold acclimation conditions for 24 h, and then returned to normal growth conditions. Isolation of sfr2 T-DNA mutants and complementation of sfi-2-3 with SFR2 cDNA. The sfi2-3 and sfr2-4 T-DNA lines (SALK_106253 and SALK_112965, respectively) were obtained from the Arabidopsis Biological Resource Center (Ohio State University, Columbus, OH). Homozygous T-DNA insertion mutants were identified by PCR-based genotyping using the following primers: sfr2-3 F P, 5’-GAGCTTGATGT'I‘CTGGAACT- 3’; sfr2-3 RP 5’-TTGAACTTTGAGCTGTCG-3’; sfr2-4 FP, 5’- CACCAAGATTGT'TGATGCATG-3’; sfi2-3 RP, 5’- TTAGTCCAGCACCACACACTG-3’. Amplification with these primers indicates the presence of wild-type genomic DNA at the respective allele; amplification with the sfr2-3 or sfr2-4 RP primers and the T-DNA left border primer LBb1.3, 5’- ATTTTGCCGATTTCGGAAC-3 ’, indicates the presence of the respective T-DNA insertion. To genetically complement sfi'2-3, the SFR2 open reading fiame was amplified 73 from cDNA by PCR using gene specific primers 5’- AAAAAGCAGGCTTCATGGAAT'TATTCGCATTGT'T-3 ’ and 5 ’- AGAAAGCTGGGTCCTAGTCAAAGGGTGAGGCTA-3 ’. Adapter primers 5 ’- GGGGACAAGTTTGTACAAAAAAGCAGGCT-3’ and 5’GGGGACCACTTTGTACAAGAAAGCTGGGT-3’ were used for secondary PCR to introduce Gateway (Invitrogen, Carlsbad, CA) cloning sequences. The resulting PCR product was inserted into the vector pDONR zeo (Invitrogen) and confirmed by sequencing. After confirming the sequence, the insert was sub-cloned into the Gateway binary vector pMDC32 (Curtis and Grossniklaus, 2003). The resulting plasmid DNA was used to transform the sfi'2-3 T-DNA mutant by Agrobacterium floral dip method (Clough and Bent, 1998). Seeds were collected from 6 different transformed plants and T1 transgenic plants were selected in the presence of hygromycin B (20pg/ml) on MS medium containing 1% sucrose. The independent T. transgenic plants were genotyped using the following PCR primers: P2, 5’-TI‘GAACTTTGAGCTGTCG-3’, and LBb1.3 to confirm the presence of sfr2-3 T-DNA and P1, 5’- CACCATGGAATTATTCGCATTGTTAA-3’, and P3, 5’- TAGAGGGTGGCTTATCAGCTGC-3’, to simultaneously confirm the absence of wild- type genomic DNA at the sfi2-3 allele and presence of transgenic SFR2 cDNA. After confirming the genotype, the leaves from independent T1 transgenic plants were phenotyped for oligogalactolipid accumulation as described below. Lipid analysis. Lipids were extracted from fresh or liquid N2-frozen MS-agar-grown plant tissue as previously described (Dormann et al., 1995). Lipids were separated by thin-layer chromatography (TLC) and visualized with H2SO4 by charring for total lipid 74 staining, or glycolipid staining with u-naphthol (Benning et al., 1995). Quantification of lipids by gas chromatographic (GC) analysis of derived fatty acid methyl esters was performed as previously described (Lu et al., 2008). Electrospray ionization mass spectrometry (ESl-MS/MS) was used as previously described (Bates et al., 2009) to analyze triacylglycerol (TAG) molecular species isolated according to (Xu et al., 2005). For 1H NMR analyses, 0.5 — 1.0 mg of DGDG was isolated from Arabidopsis leaf tissue or a scaled-up in vitro GGGT reaction (see below) by TLC. Lipids were eluted into CHC13:methanol (2:1 by ml), and were subsequently dried under N2 gas stream. Lipids were re-dissolved in CDC13:CD3OD:C2D500D (10:10:1 by vol.) and analyzed on a Varian-SOO spectrometer at 500 MHz as previously described (Xu et al., 2003). Chemical shifts were measured in relation to residual CHCl3, and data was analyzed using Mnova software (Mestralab Research, Escondido, CA). Heterologous expression of SFR2 in yeast. The Open reading frame of SFR2 was amplified by RT-PCR using Phusion polymerase (New England Biolabs, Beverly, MA) with the primers 5’-CACCATGGAATTATTCGCATTGTTAA-3’ (SFR2-F) and 5’- TCAATQATQATQATQATGATQGCCGTCAAAGGGTGAGGCTAAAG-3’ (SFR2- R); the underlined sequence indicates the coding sequence for a 6 x His-tag added directly to the C-tenninus of the SFR2 protein. The amplified product was cloned into the pYESZ. IN 5 His TOPO vector (Invitrogen) to create the vector pYESZ. 1 -SF R2, which was verified by sequencing. Saccharamyces cerevisae (Strain SCY62; genotype: MA Ta ADE2 (Sandager et al., 2002)) was transformed with pYESZ. l -SFR2 or the corresponding negative control (pYESZ). Expression of SFR2 was induced by inoculating a culture of SC minimal medium containing 2% (w/v) galactose, and 0.5 % 75 (w/v) raffinose to 0D,,00 0.4, followed by continued growth for 8 h at 28 °C. Cells were pelleted and frozen in liquid N2 and stored at -80 °C. Membranes were isolated from these cells according to (Dahlqvist et al., 2000) and stored at -80 °C. Protein quantification and immunoblotting. Membrane protein content was assayed using the RCDC kit (Biorad, Hercules, CA). For immunoblotting, samples were denatured according to (Martinez-Garcia et al., 1999), and separated on 12 % SDS- PAGE gels. Proteins were transferred to PVDF membranes and the recombinant C - terminally His6-tagged SFR2 protein was detected with anti His(C-term) monoclonal antibody (R930-25, Invitrogen) using the CPS chemiluminescent detection kit (Sigrna- Aldrich, St. Louis, MO). In vitro GGGT assay. The assay was modified from (Heemskerk et al., 1983) as follows: monogalactosyldiacylglycerol (MGDG) from Larodan (Malmo, Sweden) was dried under a N2 gas stream (64 nmol per reaction), and overlayed with 150p] per reaction assay buffer (10 mM Tricine, adjusted to pH 7.2 with Tris, 10 mM MgCl2, 0.426 mM sodium deoxycholate), which results in a final MGDszeoxycholate molar ratio of 1:2 . The reaction components were dispersed by sonication 3 x 10 sec with a Sonicator 3000 (Misonix, F armingdale, NY) equipped with a 1.6 mm diameter microtip at power setting 1.0. The reaction was initiated by the addition of 15 pg protein equivalents of isolated membranes (above) followed by sonication 1 x 10 sec as above. Reactions were stopped by the addition of 600 pl methanol:CHCl3 (1 :2 by vol.). After phase separation by addition of 600 pl 1 M KCl, 0.2 M H3PO4, and re-extraction with chloroform, lipids were separated by TLC as above. For radiometric assays, [U-3H-Gal]-MGDG, specifically labeled on the headgroup was biosynthesized using intact spinach 76 chloroplasts as previously described (Heemskerk et al., 1983). The specific activity of [U-3H-Gal]-MGDG was determined by GC and scintillation counting, and assays were performed exactly as above with 1.2 x 104 DPM [U-3H-Gal]-MGDG. Results Development of a TLC-based screen to identify Arabidopsis mutants defective in GGGT activity. Since the diagnostic products of GGGT activity, TGDG and TeDG, are not constitutively produced in wild-type plants, a treatment which activates this enzyme in viva was sought out. Earlier biochemical studies suggested increasing amounts of MgCl2 activate GGGT activity in chloroplast envelope preps (Heemskerk et al., 1983, Sakaki et al., 1990a). A series of MgCl2 solutions of increasing concentration (0-400 mM) were used to treat Arabidopsis leaves by submergence, followed by vacuum infiltration and 16-24 h of continued incubation under light. As seen in Figure 3.1, treatment with M gCl2 solutions above 100 mM resulted in the accumulation of oligogalactolipids, and 400 mM MgC 12 was used for routine screening of an EMS mutagenized Arabidopsis population. In ~1500 individual plants analyzed in a collaborative work with Bagyalakshmi Muthan, a lab technician, and an undergraduate student, Jason Sherbarth, one putative mutant defective in oligogalactolipid accumulation was recovered. To our misfortune, this mutant failed to develop and make any seeds. However, before a second isolate of this putative mutant could be recovered, an alternative approach using the same treatment to reverse-genetically screen Arabidopsis T-DNA lines yielded a candidate GGGT encoding gene, as further described below. 77 [M9051 (mM) 0 25 100 200 400 MGDG DGDG 'TGDG TeDG Figure 3.1. Activation of GGGT activity in Ieaftissue by vacuum infiltration of MgClz. Galactolipids stained with a-naphthol after separation by TLC. Wild-type leaf samples were treated by vacuum infiltration in solutions containing the indicated concentration of MgCl2, followed by 24 h incubation. 78 T-DN A mutants in the SFR2 gene are defective in oligogalactolipid accumulation. Lines for a reverse-genetics approach to identify the GGGT encoding gene were selected through two criteria: (i) the encoded protein should be present in the available chloroplast envelope proteomics data sets (Brautigam et al., 2008, F erro et al., 2002, Ferro et al., 2003, Froehlich et al., 2003)—particularly in the outer envelope where GGGT activity is unambiguously localized (Dome et al., 1982), and (ii), the encoded protein should be a member of an enzyme family involved in glycosyl transfer (either glycosyl hydrolases or glycosyltransferases) (Heinz, 1996). For the outer envelope localized glycosyl hydrolase family 1 (GH-l) encoding gene, SENSITIVE T 0 FREEZING 2 (SFR2), two homozygous T-DNA lines, sfr2-3 and sfr2-4, were confirmed by PCR (Figure 3.2). When screened using the MgCl2 infiltration assay, neither of the sfr2 mutants were able to accumulate oligogalactolipids, whereas the dgdl and dgd2 mutants defective in UDP-Gal dependent DGDG biosynthesis were similar to wild-type (Figure 3.3). The sfr2 mutants show defects in lipid remodeling during freezing stress. SFR2 was previously identified by other researchers in Arabidopsis, when null mutations in this gene were found to result in freezing sensitivity (Thorlby et al., 2004); however, the substrates and products of the encoded glycosyl hydrolase enzyme remained unknown, as did the precise mechanism through which SFR2 activity contributed to freezing tolerance (Fourrier et al., 2008). As seen in figure 3.4, the sfr2 mutant alleles exhibited freezing sensitivity similar to the previously reported alleles (McKown et al., 1996, Thorlby et al., 1999, Thorlby et al., 2004, Warren et al., 1996). Importantly, freeze-treated wild-type 79 Figure 3.2. Genotypic confirmation of the sfr2 T-DNA mutant alleles used in this study. (A) Schematic diagram of the SFR2 Gene. Exon (boxes) and intron (lines) regions are indicated as well as the open reading frame (black bkgd.) and 5’- and 3’-UTRs (white bkgd). The positions of the T-DNA insertions of the sfr2-3 and -4 alleles are indicated, as well as other mutant alleles used in other studies (sfr2-I and sfr2-2, first described by Thorlby et al., 2004 as sfiZ-I and sfi2-i2, respectively). PCR based genotyping to confirm the homozygosity of sfr2-3 (B) and sfr2-4 (C) alleles. PCR products were separated on 1% agarose gels after amplification with gene allele specific primers (FP and RP), or a T-DNA left border and complimentary gene allele specific primer (LB and RP). 80 1Kb B WT sfr2-3 WT sfr2-3 sfr2-3 FP/RP sfr2-3 LBIRP 0 WT sfr2-4 wr sfr2-4 sfr2-4 FP/RP sfr2-4 LBIRP 81 no treatment M9012 Treatment r eff is" 30" 30" a" s e6“ at is” a" c8 ’” MGDG _' TGDG I. A . > - .- quiz-L . 3g. , . ‘ .. F. . bay—:1! - " 13.- 4 - I I ~ ' V ‘ ' _‘. i U ',l‘5,§72~{~3 31", 3; [”3331 ' as: .. ‘5 . ~-‘. . ... L: >_.. :5 .- 11‘ -:. - - ‘.' ' ,- . - - .- -7 flag 3 -;~-.-. :44 26,-! _'.~ , ‘j'L. ,1 s -4__ ', ,__-- -_ ' < ‘3 .JJ‘ Figure 3.3 Accumulation of oligogalactolipids in galactolipid biosynthetic mutants. Lipids were separated by TLC and stained with a-naphtbol after extraction from fresh leaves, or leaves submerged in 400 mM MgC12, vacuum infiltrated for 5 min, and incubated for 16 h. Gene names and corresponding gene loci from The Arabidopsis Information Resource (http://www.arabidopsis.org) for the included mutants are: dgd1, Digalactosyldiacylglycerol synthase I (AT3611670); dgd2, Digalactosyldiacylglycerol synthase 2 (AT4G00550); s 2-3 and sfi'2-4, Sensitive T a Freezing 2 (ATBGO6510); tgd] -I , T rigalactasyldiacylgl ycerol I (AT 1G19800). 82 Figure 3.4. Freeze-induced galatolipid remodeling is not observed in sin mutants. (A) Appearance of wild-type (C012) and sfr2 mutants at 5 d post freezing treatment. (B) Thin layer chromatogram of lipid extracts stained for glycolipids with a-naphthol reagent from cold-acclimated (CA) or freeze-treated (FT) wild-type (WT) and sfr2 plants. (C) Changes in galactolipid levels in plants treated as in (B). ‘P < 0.05 or "P<0.01 vs. WT CA levels for 3 biological repeats. 83 B WT sfr2-3 sfr2-4 C mol % polar lipids 2.01.51.o 0.5 0 0 1‘0 20 so 49 so A leaves accumulated the oligogalactolipids TGDG and TeDG with a concomitant decrease in MGDG, while the sfr2 mutants did not (Figure 3.4B, C). Accumulation of triacylglycerols has been reported to coincide with the activation of GGGT in both the above-mentioned tgd mutants and in ozone-treated spinach (Awai et al., 2006, Lu et al., 2007, Sakaki et al., 199%, Sakaki et al., 1990c, Xu et al., 2005, Xu et al., 2008). Indeed, TAG was found to accumulate to ~6% of total fatty acids as a result of freezing treatment when compared to cold acclimated wild-type plants (Figure 3.5A, B). The sfr2 mutants showed defect in TAG accumulation and a large decrease in 16:3 (carbonzdouble bonds) acyl groups esterified to TAG in freeze-treated plants compared to wild type (Figure 3.5C). The presence of 16:3 containing TAG species in wild-type leaves was confirmed by mass spectrometry (Figure 3.6). The 16:3 acyl group is predominately found esterified to MGDG in Arabidopsis (Browse et al., 1989), indicating that DAG produced by GGGT activity is, in part, frnther acylated to TAG. Effect of various salts and osmolytes on oligogalactolipid and TAG accumulation following vacuum infiltration. To test whether there was any specificity for Mng in activating GGGT by vacuum infiltration, a variety of other salts were tested for their ability to result in the accumulation of TGDG (Figure 3.7A) or TAG (Figure 3.73). All salts tested (MgSO4, CaCl2, BaCl2, NaCl, and NH4Cl—all at 0.4 M) activated TGDG and TAG accumulation. These results indicated that neither the identity nor the ionization potential of the cation or anion in the salt were specifically required. To test whether the high ionic strength present in the various salts tested, in general, was required for TGDG and TAG accumulation, various concentrations of the non-ionic osmolytes glycerol and 85 Figure 3.5. Freeze-induced Triacylglycerol accumulation. (A) Thin layer chromatogram of neutral lipids visualized by H2SO4 and charting fi'om wild-type (WT) and sfr2 plants treated as in Figure 3.4. (B) Percent of total fatty acids esterified to TAG in plants treated as in (A); the average and standard deviation of at least three biological repeats are shown. (C) Fatty acid profile of TAGs in plants nested as in (B) shown on a mol % basis of the average and standard deviation of at least three biological repeats; "P < 0.01 or """"P<10'3 vs. WT FT levels. The labels along the X-axis indicate the fatty acid species identified (number of carbonsznumber of double bonds). 86 A WT sfr2-3 sfr2-4 CA FTFTFT TAG- FFAs- DAG- B 8 . (I) . :9 7 s 6‘ E 5 ' a 4- (D 3 . < P 2 I s 1 , 0 . WT C CA 60' nwr CA 50‘ IWI'FT esfi2-3 FT 40‘ nsfi2-4 FT 101 Fatty acid profile (mol %) 00 9 O . u 'H H 16:0 16:1 16:3 18:0 18:1 18:2 18:3 wr sfr2-3 sfr2-4 FT FT FT 87 > 52 x 541x 1001 ‘ ' 8 C (B D C 3 A it e". 0 > '5 g i 1111 0 1 ._, Ill vllr lllerJn ‘ illi llllllll. ‘ lllllu. ' Y. . Y 1 In #I. 7 ' 1.1 .‘l T I 800 820 840 860 880 900 920 940 960 980 1000 B mlz 100. / lM+NH} NH}18:3C Rcoo'l Daughter ions of 863 l / {mun} NH}16 3c RCOO] Relative Abundance (°/o) 0-11 . -- 540 550 560 570 580 590 600 610 620 630 640 650 660 670 680 690 C mlz [M+NH} NH}18:2C Rcoo'] Daughter ions of 865 100' \[Mi-NH} NH}18:3C Flood] 32 ~ [M+NH} NH}16:30 R000] 8 fl 3 .0 2‘. g .9 0 a: 540 550 660 570 580 590 600 610 620 630 640 650 660 670 680 690 mlz Figure 3.6. ESI-MS/MS of triacylglycerols isolated from freeze-treated wild-type plants. (A) ESI-MS spectra of TAGs isolated from fi'eeze treated wild type leaves. The numbers above the brackets indicate the number of carbons present in the three acyl chains; x denotes the variation in degree of unsaturation of the acyl chains. ESI-MS/MS spectra of the daughter ions of parent ions 863 m/z (B) and 865 (C). Deduced identities of the daughter ions are indicated in brackets. 88 Figure 3.7. Salt and osmolyte treatments that activate TDGD and TAG accumulation in wild-type leaves. a-naphthol stained galactolipids (A, C) and triacylglycerols visualized by H2804 charring (B, D) after separation by TLC from leaves treated by vacuum infiltration with water (H20) or the indicated salt solutions at 0.4 M (A, B), or by the osmolytes glycerol and sorbitol (C, D). Vacuum infiltration was for 5 min, followed by 16 h incubation before lipid extraction; 4 M-V indicates that vacuum infiltration was not used, and the tissue was only submerged in the given osmolyte at 4 molar. In (C) and (D), the numbers indicate the molarity of the osmolyte, and water and MgCl2 controls as treated in (A) were included. 89 >. 2 V 0 Sorbitol 20410204 0204102040v'0 sorbitol were employed in the vacuum infiltration assay. As seen in Figure 3.7C-D, use of solutions at a concentration 2 1.0 M of the osmolyte were sufficient to cause TGDG and TAG accumulation, and at 4.0 M, vacuum infiltration was unnecessary for this effect. These results indicated high extracellular osmotic pressure is the underlying factor in the activation of GGGT in planta by this method. Genetic complementation of sfr2-3 by transgenic over-expression of the SFR2 cDNA. To confirm that the T-DNA insertion in sfi2—3 is causing the oligogalactolipid accumulation defect described above (Figures 3.3 and 3.4), the SF R2 cDNA was cloned from wild-type tissue, and inserted into the constitutive expression vector pMDC32 (Figure 3.8) as described in “Methods”. The sfr2-3 mutant plants were transformed with this plasmid, and the resulting T1 progeny for seven individual transformants were found to accumulate oligogalactolipids when tested with the MgCl2 infiltration assay (Figure 3.8B-C). These results clearly indicated that the presence of transgenic, wild-type SF R2 cDNA was sufficient to complement the TDGD accumulation defect phenotype of sfi'2-3. Heterologous expression of the SFR2 protein in yeast and analysis of in vitro GGGT activity. The SFR2 protein was produced in yeast by expression of the SFR2 cDNA and localized to the isolated membrane fraction by immunoblotting SDS-PAGE separated protein preparations with an antibody that recognizes a c-terminal 6 x Histidine tag added to SFR2 during cloning (Figure 3.9A). The protein:fatty acid ratios (ug nmol") of membranes isolated from empty vector and SF R2-expressing strains were similar (Figure 3.98). Incubation of SFR2-containing membranes with deoxycholate dispersed MGDG 91 Figure 3.8. Complementation of the sfi-2-3 T-DNA mutant with SFR2 cDNA. (A) Schematic diagrams of the A. thaliana chromosome 3 (AtChr3) structure of the 5’ end of the SFR2 encoding gene and the SFR2 cDNA expressing transgenic insertion derived from the plasmid pMDC32 +SFR2. Positions of primers used in genotyping (B) are indicated. (B) Genotyping of C012, sfr2-3 and sfr2-3 :: SFR2 complementing lines with primers that specifically detect the sfr2-3 T-DNA insert (LBb and P2), and primers which detect the presence of a wild-type genomic copy of SFR2 (638 bp), or the presence of transgenic SF R2 cDNA (402 bp). (C) Phenotypic complementation of sfi'2-3 oligogalactolipid accumulation defect in the sfi2-3::SFR2 lines. leaves from plants were vacuum infiltrated with 400 MgCl2 and incubated for 16 h, followed by separation of extracted lipids by TLC and visualization of galactolipids (MGDG, DGDG, TGDG and TeDG) by staining with a-naphthol. Only the 7 complementing lines genotyped in (B), analyzed on two separate TLC plates are shown, along with wild-type and sfi2-3 controls. 92 //L“ 2x 358 ms 577 +SFR2 B .5 sfr2-3 :: SFR2 db ‘§L 1 2 3 4 5 6 7 LB + P2 0.. P1 (I'll-DII'IIIH-Illliilllas (3 PER” meez m + - --~ ..... l-I‘ .' -*0 a . ‘17 MGDG "'7’” “‘wrfm 1:11:— .‘ ,. ,1. DGDG1"1!H!5'!E!!!P .p-I-IinIUIQp» TGDG " i , TBGDG ref . w- m “P 1 2 3 ‘1’ 4 5 6 7 E % sfr2-3 :: SFR2 E % sfr2-3 :: SFR2 93 A A N N c or 'o '01 l l l I pg protein nmol'1 fatty acid 0 'ur O I EV +SFR2 Figure 3.9. Heterologous expression of SFR2 in Saccharamyces cerevisae. (A) Anti His (C-term) antibody immunoblot (upper panel) and Coomassie Brilliant Blue stained gel (lower panel) of SDS-PAGE separated membrane protein preparations (25 pg protein) for empty vector control (EV memb.) and SFR2 expressing (+SFR2 memb.) yeast strains; soluble proteins fiom the SFR2 expressing strain (+SF R2 sol.) were also included. The positions of a molecular weight standard are indicated in kDa. (B) Analysis isolated membrane fractions for protein content (ug) normalized on a per nmol total fatty acid basis for the empty vector and SFR2 expressing strains as described in (A). 94 resulted in the production of DGDG, TGDG, and TeGDG at l h incubation, whereas empty vector control membranes did not (Figure 3.10A). In a time course using [U-3H- Gal]-MGDG, SFR2-containing membranes converted ~60% of labeled MGDG into DGDG and oligogalactolipid products in 3 h. This conversion was markedly reduced by the addition of EDTA equivalent to Mg2+ ions present (10 mM) (Figure 3.10B-C) as was previously observed in chloroplast envelopes (Heemskerk et al., 1983). The fatty acids 16:3 and 18:3, specific to the plant derived MGDG added to the assay, were present in the DAG pool only in the SF R2 protein-containing reactions—indicating DAG is being produced from MGDG by SFR2 in vitro (Figure 3.10D). Proton-NMR spectra of DGDG isolated from a scaled-up reaction showed the glycosidic linkages to be all in the fl-nomeric configuration (flflDGDG), which is distinct from the flaDGDG synthesized by the UDP-Gal dependent DGDI/2 enzymes found in wild-type leaves during normal grth (Benning and Ohta, 2005, Xu et al. , 2003) (Figure 3.11). The retention of configuration from fiMGDG to fiflDGDG by SFR2 is consistent with all other GH-l enzymes, which are classified as retaining fl-glycosidases with a broad range of substrate specificities (Hill and Reilly, 2008). Discussion In this work, a longstanding unknown enzyme in chloroplast galactolipid metabolism, the galactolipid:galactolipid galactosyltransferase (GGGT) has been identified, and is encoded by the previously identified gene of unknown function, SFR2 (Thorlby et al., 2004). Clearly, GGGT is not essential for normal growth and development, but rather for sub-zero temperature lipid remodeling of the chloroplast 95 Figure 3.10. Recombinant SFR2 in vitro activity from transgenic yeast. (A) Thin layer chromatogram of lipid extracts stained for galactolipids with a-naphthol reagent from in vitro GGGT activity assays using deoxycholate/MGDG dispersions incubated with membrane fractions from empty vector control (EV) or SFR2 expression strain (SFR2). The in vitro reactions were stopped after 1 h of incubation. Lipids from the reactions were extracted, chromatographed and compared to lipids from a freeze-treated leaf sample (FT Leaf). (B) In vitro GGGT activity using labeled [U-3H-Gal]-MGDG. The data are presented as the percent of 3 H label found in DGDG, TGDG and TeDG over time. Empty vector and SFR2 were assayed as in (A), and a SFR2 membrane protein assay containing EDTA at a concentration equimolar to Mg2+ present (10 mM) is also shown. Data are representative of two measurements, and the standard deviation is shown if it exceeds symbol size. (C) Percent of [3H] label found in different galactolipid species in the SFR2 assay shown in B. (D) Fatty acid composition of diacylglycerol in empty vector control (EV) and SF R2 reactions at 3 b. Data are presented on 3 mol% basis as the average and standard deviation of three biological repeats. 96 C100 % label in lipid species % [3H]MGDG converted m -o- MGDG D *DGDG *TGDG/TeDG 30 60 90 120150180 time (min) 97 80‘ 60- $ 0" O O a a O a DAG fatty acids (mol%) 8 0) _a C I O a +SFR2 *SFRZ + EDTA q—EV 30 60 90 120150180 time (min) I EV a SFR2 16:016:1 16:318201821 18:3 fatty acid species Figure 3.11. IHNMR spectra of galactolipids from plant leaves and isolated from the in vitro GGGT assay. MGDG (A) from the commercial supplier Laurodan, DGDG (B) isolated from wild-type (C012) leaves, and (C) DGDG isolated from a scaled-up in vitro GGGT reaction. 98 H1’ H1a 1b 11 1 11 LL __ ,‘~ 1 ‘ am“ 1’ __ :w~- J“ M -2 All” I &l 4] 4b ' 45 44 43 . 4h . 4H 40 3b ' as 41 1 4b 4b ' 4h 43 ' 42 ' 4H ' 4o . 33 . 33 ppm 99 envelope during freezing stress. Through consideration of (i) what is currently known about plant cell responses to freezing, (ii) the work conducted by others on the sfr2 mutants in Arabidopsis prior to the functional elucidation of SFR2, and (iii) the work presented above, a model for the role of GGGT in freezing tolerance can be envisioned. A hypothesis for the role of GGGT in stabilizing chloroplast envelopes during freeze-induced cellular dehydration. In order to understand how GGGT contributes to freezing tolerance in plants, one must first consider the wealth of physiological, biochemical, and molecular/ genetic insight that has been established in this intensely studied field. Freezing damage is primarily visited upon the cell membranes and is attributed to severe cellular dehydration, which is initiated by extracellular ice formation that results in decreased water potential across the plasma membrane (Steponkus, 1984, Thomashow, 1999). Under these conditions the formation of non-lamellar lipid phases, such as the inverted hexagonal II (H11) phase can occur and, thus, stabilization of lamellar membrane systems within the dehydrated plant cell is a key determining factor in the survival of freezing (Steponkus, 1984, Thomashow, I999, Uemura et al., 1995). To survive mild freezing, plants like Arabidopsis require pre-exposure to low, non-lethal temperatures in a process known as cold acclimation (Hirayarna and Shinozaki, 2010, Thomashow, 1999, Uemura et al., 1995). Cold acclimation involves transcriptional (Guy et al., 1985, Kilian et al., 2007, Zeller et al., 2009) and metabolic (Cook et al., 2004, Kaplan et al., 2004) changes which result in multiple mechanisms protecting against freezing damage—including increases in intracellular solutes and the accumulation of cryoprotective metabolites (Uemura et al., 100 2003) and proteins (Steponkus et al., 1998). The phenotype of Arabidopsis plants carrying mutations in the SENSITIVE T 0 FREEZING 2 (SF R2) gene, however, clearly indicates a distinct mechanism required for freezing tolerance (Fourrier et al., 2008, McKown et al., 1996, Thorlby et al., 2004, Warren et a]. , I996). The sfrZ mutants show extensive intracellular damage following freezing recovery, with rupture of both chloroplasts and tonoplasts—likely through fusion of destabilized membranes in these organelles. The SFR2 protein is constitutively present, and sfr2 mutants are not affected in the regulation of known cold-acclimation processes. With cellular membrane damage in mind, alterations in lipid composition during cold acclimation, such as increased fatty acid unsaturation and phospholipid content, have long been correlated with enhanced freezing tolerance, but are not yet distinguished from adaptations that arise from growth at low above-freezing temperatures rather than freezing stress per se (Nishida and Murata, 1996, Steponkus, 1984, Thomashow, 1999). More recent lipid profiling studies in Arabidopsis have revealed that more drastic lipid compositional changes occur during freezing, including a decrease in the chloroplast- specific lipid monogalactosyldiacylglycerol (MGDG) (Du et al., 2010, Li et al. , 2008, Welti et al., 2002). Our hypothesis for the requirement of SFR2-dependent galactolipid remodeling in freezing tolerance centers on the prevention of non-lamellar Hn phase formation brought about by dehydration (Figure 3.12). During dehydration, Hn phase is formed at the interface of apposed bilayers, and believed to initiate at the chloroplast envelope during freezing (Steponkus, 1984, Steponkus et al., 1998, Thomashow, 1999). This results in fusion between bilayers (Siegel and Epand, 1997), particularly when membranes are 101 sfr2 SFR2 ,. ,“t r. trtrt rutl'Hl the CM ' l)--t..l rlrlll'l/lllllt I'll .. r " 3DAG —> STAG J MGDG ‘4 teens MGDG —> TGDG ZMGDG —" DGDG All I ;‘ )kéxl‘oI/I. .. 1.1)-71”,“ I'llf mm 1 H "‘ll"-l'lll.t"l‘ troll/.74?! luVrrlirrllr-ali-YI‘ OEV Figure 3.12. Model of SFR2 activity and the effect of freezing on the chloroplast outer envelope membrane (eEv) and adjacent cell membranes (CM) in the sfi'z mutant and the wild type (SFR2). Lipid bilayers and hexagonal 11 phase (113;) formation leading to membrane fusions in sfr2 following freezing are indicated. SFR2 is shown associated with the outer leaflet of the outer chloroplast envelope membrane. Three cycles of the reaction are indicated consuming 4 moles of MGDG and producing 1 mole of TeGDG and 3 moles of TAG. This assumes that every mole of DAG produced is converted to TAG which may not be the case as some of the DAG might be converted to phosphatidic acid. 102 enriched in glycerolipid species with relatively small head groups (e. g. MGDG and phosphatidylethanolamine) as these show a higher propensity for transition to H1] phase. Previously, sfr2 mutants showed extensive chloroplast and tonoplast rupture in leaves during freezing recovery, which was proposed to arise from fusion of destabilized membranes (Fourrier et al., 2008). This work shows that SFR2 partially converts MGDG to DGDG and oligogalactolipids (GGGT activity) which are not prone to form Hn phases. This is analogous to the UDP-Glc-dependent modulation of mono- to di-glucolipid ratios observed in Acheloplasma laidlawii under different abiotic stress conditions (W ieslander er al., 1986). One distinction is that DAG is produced by SFR2, and is further metabolized to TAG, and possibly other lipid species, as accumulation of DAG, which can form non-lamellar phases, must be prevented. Moreover, the action of SFR2 provides a mechanism by which membrane lipids and excess membrane are removed to accormnodate a shrinking organelle following freezing. Under the conditions of fieeze- induced dehydration, a response initiated by transcriptional activation may not be practical for reasons including the time required for transcription, translation, protein folding and lastly, localization, and also that these processes may be defective under this stress. As such, the constitutive presence of inactive GGGT on the chloroplast envelope, which could be directly activated by sensing physical changes in the membrane during freezing, may offer a more successful strategy in mitigating membrane damage. Evolution of glycosyl hydrolase family enzymes towards catalysis of transglycosidation: a proposed mechanism for GGGT activity. Many GH-l family enzymes are known to catalyze moderate levels of transglycosidase activity under certain 103 in vitro conditions, and examples for which transglycosylation predominates over hydrolysis are known in other GH families (e. g. GH-70 glucansucrases) (Sinnott, 1990). SFR2, which was found to be most divergent among 253 GH-l family proteins by phylogenetic analysis (Marques et al., 2003), has evolved to catalyze MGDG transgalactosidation (GGGT activity) by exclusion of water as an acceptor—which could be attributed to several intrinsic features of SFR2—including the exclusory binding of acceptor galactolipids, and/or the close apposition of the catalytic domain to the membrane surface. As seen in figure 3.13, the proposed GGGT mechanism is a modified version of the classical Koshland double displacement mechanism (Koshland, 1953), where the enzyme-linked galactoside intermediate from donor MGDG is transferred to MGDG (or DGDG/TGDG through processive activity) rather than water as an acceptor. As such, SFR2 may provide insight into improving the transglycosidase activity of other GH-l enzymes to produce oligosaccharides of medical or industrial importance (Perugino et al., 2004). Is the protective role of GGGT specific to freeze-induced dehydration? Orthologs of SFR2 are found in all land plants with completed genomes (Fourrier et al., 2008). Intriguingly, SFR2 is also conserved in species with no tolerance to freezing (e.g. tomato, maize and rice) (Figure 3.14), suggesting SFR2 function is not restricted to freezing protection (Fourrier er al., 2008). There is a large overlap in the mechanisms required for freezing tolerance and dehydration due to water deficit or high salinity (Hirayama and Shinozaki, 2010, Thomashow, 1999), and SFR2 orthologs in these species may act on membrane stabilization during other abiotic stresses which cause cellular dehydration. 104 Figure 3.13. A proposed reaction mechanism for GGGT. The reaction is initiated MGDG substrate binding, followed by nucleophilic attack by an acidic (Glu) residue (step 1), which is assisted by another Glu residue acting as an acid catalyst in forming the first transition state (step 2). Cleavage of the glycosidic bond results in the release of the aglycon moiety (RIOH, diacylglycerol [DAG]), and the formation of a glycosyl enzyme intermediate (step 3). The glycosyl donor substrate (R20H, MGDG) binds and is deprotonated by the Glu residue, now acting as a base catalyst, followed by attack of the enzyme-linked glycoside to form the second transition state (step 4). The enzyme linked glycosidic bond is cleaved, resulting in the transglycosidation of bound MGDG to form BBDGDG (step 5). The BBDGDG product is released, or' through processive activity, can be further transgalactosylated (via steps 1-5) forming TGDG and subsequently TeDG. The steps are delineated by numbered grey boxes. 105 0-05 r——-P. patens 1 100 P. patens 2 P. tn'chocarpa 100 100 R. comums ~ V. vinifera 64 S. chopersicum G. max 2. mays .l 68 O. sativa (lndica) 100 79 ———- B. distach yon A. thaliana (SFR2) P. teada Figure 3.14. Phylogenetic relationships of SFR2 orthologs. Bootstraping values (1000 replicates) are shown at the nodes as percentages. The Genbank accession numbers for the amino acid sequences are: Arabidopsis thaliana (SFR2), CAD36513; Brachypodium distachyon, (ACF 22735); Glycine max, (CAJ 87636); Oryza sativa-Indica, (EAY81793); Physcometrilla palens l, (XP_001759357); P. patens 2, (XP_001756957); Pinus teada, (CAJ 87639); Populus trichocarpa, (XP_002316058); Ricinus communis, (XPOOZS 19275); Solanum chopersicum, (CAJ87637); Vitis vinifera, (C8122845); Zea mays, (NP_001105868). 107 Infiltration of Arabidopsis leaves with any osmotically active compound tested induced GGGT activity (Figures 3.1 and 3.7). While the sensitivity of sfl2 mutants to severe drought remains to be tested in detail, a number of physiological studies in crop species (e.g., wheat and maize) and dessication tolerant resurrection plants (e. g., Ramonda serbica), have found decreases in MGDGzDGDG ratios and the accumulation of TAG in response to water deficit treatments (Navari-Izzo and Rascio, 1999). Whether SFR2 orthologs are participating in dehydration-induced lipid remodeling events in these species is currently unknown. Regardless, it is emerging that modulation of membrane lipid composition in response to cellular dehydration is a critical factor in the survival of this stress. Future study of SFR2 may also provide general insight as to how organisms outside the plant kingdom, including prokaryotes, yeasts, and some animals have evolved tolerance to severe dehydration—and even desiccation—which was likely an important factor in the adaptive transition of aquatic species to life on land. References Awai, K., Xu, 0, Tamot, B., and Benning, C. (2006) A phosphatidic acid-binding protein of the chloroplast inner envelope membrane involved in lipid trafficking. Proc. Natl. Acad. Sci. U. S. A 103, [0817-1082. Bates, P. D., Durrett, T. P., Ohlrogge, J. B., and Pollard, M. 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Plant J. 58, 1068-1082. 113 Chapter 4 Molecular genetics of lipid metabolism in the model green alga Chlarnydomonas reinharda'r’ 3 Parts of this work were published in: Moellering, E. R., Miller, R., and C. Benning. "Molecular Genetics of Lipid Metabolism in the Model Green Alga C hlamydomonas reinhardtii" Lipids in Photosynthesis: Essential and Regulatory Functions. H. Wada and N. Murata, eds, Springer Netherlands-Dordrecht, 2009. Pp. 139-155. 114 Abstract Lipid biosynthesis in algae has gained renewed attention in recent years, as algal lipids are increasingly seen as promising biofuel and nutraceutical feedstocks. In contrast to the molecular-genetic insight in seed plant glycerolipid metabolism, the genetic factors that govern lipid biosynthesis in the diverse group of species collectively referred to as algae are less well known. While the unicellular green alga Chlamydomonas reinhardtii is unlikely to be developed for commercial production, it is currently the best genetic and genomic model for microalgal lipid research. This chapter focuses on the current knowledge of lipid metabolism in this alga, as well as the accumulation of neutral glycerolipids (e.g., triacylglycerols [TAGs]) in different algal species during environmental stress. Distinctions between seed plant and Chlarnydomonas lipid metabolism are also discussed. For example, Chlarnydomonas lacks phosphatidylcholine and has in its place the betaine lipid diacylglyceryl-N, N, N,-trimethylhomoserine. This has important implications for lipid trafficking and lipid modification—including the accumulation of TAGS, an increase of which is a major goal in improving algae as a biofuel feedstock. 115 Abbreviations ACP CDP-DAG DAG DGDG DGTS ER FAS PC PG PE Pl PA PS PUFA MGDG RNAi SQDG TAG acyl carrier protein CDP-diacylglycerol diacylglycerol digalactosyldiacylglycerol diacylglyceryl-N, N, N,-trimethyl homoserine endoplasmic reticulum fatty acid synthase phosphatidylcholine phosphatidylglycerol phosphatidylethanolamine phosphatidylinositol phosphatidic acid phosphatidylserine polyunsaturated fatty acid monogalactosyldiacylglycerol RNA interference sulfoquinovosyldiacylglycerol triacylglycerol 116 Introduction Glycerolipid biosynthesis in plants has been intensely studied for decades, providing substantial molecular insight into plant lipid metabolism. Through a combination of genetic and biochemical approaches, the genes encoding enzymes for glycerolipid biosynthesis and fatty acid desaturation have largely been identified (Benning and Ohta, 2005, Frentzen, 2004, Holzl and Dormann, 2007, Joyard et al., 1998, Ohlrogge and Browse, 1995), and more recently, factors involved in endoplasmic reticulum (ER)-to-plastid lipid trafficking have been discovered (Awai et al., 2006, Benning, 2008, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008). Furthermore, sequencing of plant genomes, beginning with Arabidopsis (The Arabidopsis Genome Initiative, 2001), followed by functional annotation of the encoded genes has allowed for the identification of hundreds of candidate lipid metabolism and trafficking proteins that await functional characterization (Beisson et al., 2003). Chlamydomonas reinhardtii is a well established eukaryotic green microal gal model organism, which has been utilized for the study of different processes including photosynthesis (Niyogi, 1999), post-transcriptional gene silencing (Wu-Scharf et al., 2000), cell biology of flagella (Silflow and Lefebvre, 2001), and microalgal metabolism (Grossman et al., 2007). The availability of the complete Chlarnydomonas genome sequence (Merchant et al., 2007), as well as a number of available molecular tools, including facile nuclear insertional mutagenesis (Tam and Lefebvre, 1993), RNA interference methods (Fuhrmann et al., 2001, Molnar et al., 2009), and a molecular map (Kathir et al., 2003), make C hlamydomonas a promising model organism for the discovery of genes involved in lipid metabolism and trafficking using both forward and 117 reverse genetics. Initial annotations of Chlamydomonas genes involved in lipid metabolism have recently been published (Riekhof et al., 2005b, Riekhof and Benning, 2008), which highlight both the similarities and differences in land plant and green algal glycerolipid metabolism. With the resurgence of interest in using microalgae as a biomass feedstock for renewable transportation fuels that does not have to compete with resources used in agriculture (Hu et al., 2008), the availability of a suitable model microalgal organism is timely. Even though Chlamydomonas reinhardtii is itself an unlikely candidate species for production of biofuels, it is related to other unicellular algal species (e. g. Dunalliella salina) that are used for commercial scale production of lipids (Hu et al., 2008). Chlarnydomonas has been reported to accumulate triacylglycerols (TAGS) under conditions of nutrient deprivation (Weers and Gulati, 1997) or high light (Picaud et al., 1991), which has been reported in many other green algal species (Hu et al., 2008). C hlamydomonas also synthesizes TAGS from lipids supplied in the medium (Grenier et al., 1991). As such, insight into glycerolipid and TAG metabolism gleaned from studies in Chlarnydomonas may be instructive in the rational engineering of other algal species for biofuel production. Below, the current understanding of glycerolipid biosynthesis in Chlarnydomonas and TAG metabolism in different algal species are discussed. Fatty Acid Synthesis and Incorporation into Glycerolipids. De novo synthesis of fatty acids is localized to the chloroplast of Chlamydomonas cells (Sirevag and Levine, 1972), and the common ancestral origin of green algal and seed plant plastids (Reyes-Prieto et al., 2007) is particularly apparent in many homologous components of the fatty acid biosynthetic machinery. For example, 118 bioinformatic analysis of the Chlamydomonas genome has identified genes for the full suite of enzymes required for the conversion of acetyl—CoA to acylated-acyl carrier protein (ACP), including the multimeric bacterial-type acetyl-CoA carboxylase and fatty acid synthase complexes (Riekhof et al., 2005b, Riekhof and Benning, 2008). These enzymes are essential for fatty acid biosynthesis in plants (and presumably algae) which predominantly produce 16:0-ACP and 18:1-ACP as the result of desaturation of 18:0- ACP by a soluble stearoyl-ACP A9 desaturase (Browse and Somerville, 1991, Shanklin and Somerville, 1991). As in plants, fatty acids are incorporated directly into chloroplast membrane glycerolipids in Chlarnydomonas (Figure 1) by stepwise acylation of glycerol 3-phosphate to form phosphatidic acid (snI-18zl, sn2-16:0-PA) by glycerol 3-phosphate : acyl-ACP acyltransferase (GPAT), which shows substrate specificity for 18:1-ACP, and then by lysophosphatidate : acyl-ACP acyltransferase (LPAT, 16:0-ACP specific) (Browse and Somerville, 1991, Kim and Huang, 2004, Kunst et al., 1988, Murata and Tasaka, 1997, Xu et al., 2006). Fatty acids are also assembled into glycerolipids at the ER where isofonns of the plastid acyltransferases are present and have been characterized in Arabidopsis (Kim et al., 2005, Zheng et al., 2003). Putative orthologs of the plant GPAT and LPAT genes are annotated in the final Chlarnydomonas genome draft (Riekhof et al., 2005b, Riekhof and Benning, 2008). Candidates for the plastid PA phosphatase which produces the diacylglycerol precursors for the biosynthesis of non-phosphorus lipids in the plastid has been recently identified in Arabidopsis (Nakamura et al., 2007). However, there is currently no good candidate in the Chlamydomonas genome predicted to encode this enzyme (Riekhof et al. , 2005b, Riekhof and Benning, 2008). 119 Figure 4.1 Overview of glycerolipid biosynthesis in Chlarnydomonas. End-product glycerolipids in the plastidic and endoplasmic reticulum (ER) pathways are shown in bold. Abbreviations used are: ACP, acyl carrier protein; AdoMet, S-adenosylmethionine; ASQD, 2'-O-acyl-sulfoquinovosyldiacylglycerol; CDP, cytidine-5'-diphosphate; CoA, coenzyme A; CTP, cytidine-5'-triphosphate; DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; DGTS, diacylglyceryl-N,N,N-trimethylhomoserine; Etn, ethanolamine; FA, fatty acid; G-3-P, g1ycerol-3-phosphate; Glc, glucose; Ins-3-P, inositol-3-phosphate; MGDG, monogalactosyldiacylglycerol; P-Etn, phosphoethanolamine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PGP, phosphatidylglycerolphosphate; PI, phosphatidylinositol; PA, phosphatidic acid; Ser, serine; SQ, sulfoquinovose; SQDG, sulfoquinovosyldiacylglycerol; TAG, triacylglycerol; UDP, uridine-5'-diphosphate. Key enzymatic steps (1-14, plastid; 15-23, BR) are indicated by boxed numbers next to the arrows indicating substrates and products: 1, acetyl-CoA carboxylase; 2, Malonyl-CoAzACP transacylase; 3, acyl-ACP thioesterase; 4, G-3-P acyltransferase; 5, lyso-PA acyltransferase; 6, CDP-DAG synthase; 7, PGP synthase; 8, PGP phosphatase; 9, PA phosphatase 10, sulfolipid synthase; ll, sulfolipid 2’-0-acyltransferase; 12, MGDG synthase; l3, DGDG synthase; l4, UDP-SQ synthase; 15, long chain acyl-CoA synthetase; l6, G-3-P acyltransferase; l7, lyso-PA acyltransferase; 18, PA phosphatase; 19*; ER PG synthesis pathway (same as steps 6 to 8); 20, CDP-DAGzinositol phosphotransferase; 21, DAG acyltransferase; 22, betaine lipid synthase; 23, CDP-EtnzDAG ethanolamine phosphotransferase. 120 ~, / Plastid Acetyl-CoA ASQD DGDG IT El IT UDP-Gal SQDG DAG—i MGDG Malonyl-CoA UDP-Gal UDP so n 1211”" 3032/. PA—C@—>CDP- DAG MalonyI-ACP UDP-Glc El file-w PAS Lyso-PA PGP Acyl-ACP Ii / G- - PG Free-FA 1 L r‘ifi 7 Cytoplasm/ER \T/ F FA fee. G-3-P Acyl-CoA/j- TAG El Lyso-PA Ser CDP-Etn l DAG {-— pA Etn—>P-Ern @lAdoMet P@/ Ext-P. -3-P DGTS 121 Chloroplast Membrane Lipids. The overall structural organization of membranes in the chloroplast of Chlamydomonas and seed plant chloroplasts is essentially identical, where the inner and outer envelope membranes enclose an extensive thylakoid membrane system in which the photosynthetic apparatus is embedded. Genetic studies of Arabidopsis have identified many of the genes responsible for the biosynthesis of chloroplast membrane lipids, and have revealed the essential role that lipid composition plays in optimal photosynthetic function (Benning and Ohta, 2005, Dormann and Benning, 2002, Vijayan et al., 1998, Wallis and Browse, 2002). Though the genetic study of glycerolipid metabolism in C hlamydomonas has far fewer documented examples, detailed biochemical analysis of this alga’s lipid composition has long confirmed the presence of the major chloroplast membrane lipids found in land plants—including the galactoglycerolipids mono- and di-galactosyldiacylglycerol (MGDG, DGDG), sulfoquinovosyldiacylglycerol (SQDG), and the phosphoglycerolipid phosphatidylglycerol (PG) (Giroud et al., 1988). As in plants, galactoglycerolipids are the predominant membrane glycerolipid class in Chlarnydomonas, where they make up a majority of the chloroplast membrane lipids (Giroud et al., 1988, Janero and Barnett, 1981a). In Arabidopsis, the bulk of galactolipid biosynthesis involves two enzymatic steps (Figure 1), whereby MGDG is formed from diacyl glycerol (DAG) and UDP-galactose (UDP-Gal) substrates by MGDG synthase (MGDl ), and DGDG is formed from MGDG and UDP-Gal by DGDG synthase (DGDI) (Awai et al., 2001, Benning and Ohta, 2005, Dormann et al., 1999). Genes encoding MGDG and DGDG synthases have been identified in the C hlamydomonas genome as orthologs of the Arabidopsis genes MGD] and DGD], respectively (Riekhof 122 et al., 2005b, Riekhof and Benning, 2008). MGD] and DGD] are single-copy genes in Chlamydomonas, which differs from that of the MGD], 2, 3 and DGD], 2 paralogs found in the Arabidopsis genome. Molecular analysis of the Arabidopsis MGD2, 3 and DGDZ genes has revealed their role in a galactolipid biosynthetic pathway which is transcriptionally induced during phosphate deprivation, and is proposed to provide galactolipids for extraplastidic membranes (Hartel et al., 2000, Jouhet et al., 2004, Kelly et al. , 2003, Kelly and Dormann, 2002). The apparent lack of this induced galactolipid pathway in Chlarnydomonas suggests a distinct lipid metabolic response to phosphate limitation, or a lack of need for one. However, to date the galactolipid biosynthetic genes of Chlamydomonas have not been studied in detail at the molecular level to test these hypotheses. The sulfolipid sulfoquinovosyldiacylglycerol (SQDG) has long been studied in the context of its role in photosynthetic membranes, not only due to its prevalence in photosynthetic eukaryotes and prokaryotes, but also because of its association with photosynthetic pigment-protein complexes (Gounaris and Barber, 1985, Menke et al., 1976, Pick et al., 1985, Stroebel et al., 2003). However, the more recent discovery of SQDG and the genes and enzymes involved in SQDG biosynthesis in non-photosynthetic bacteria and archea as summarized in (Benning, 2008, Cedergren and Hollingsworth, 1994), has clearly indicated that the role of sulfolipids is not limited to the function of photosynthetic membranes. The biosynthesis of SQDG is carried out in two enzymatic steps in Arabidopsis by (i) SQD] , which catalyzes the formation of UDP-sulfoquinovose from UDP-Glc and sulfite, and (ii) SQD2, which transfers the sulfoquinovose moiety from UDP-sulfoquinovose to DAG, forming SQDG (Essigmann et al., 1998, Sanda et al., 123 2001, Yu et al., 2002). A single copy ortholog of SQDI is present in Chlamydomonas, and two possible orthologs of Arabidopsis SQDZ (SQD2, 3) are found in the genome (Riekhof et al., 2003, Yu et al., 2002). Recently, a Chlarnydomonas mutant deleted in SQDI (Asqdl) and completely lacking sulfolipid has been studied (Riekhof et al., 2003). Phenotypic analysis of Asqdl revealed a reduced growth rate during phosphate-limiting conditions—under which the SQDG level was found to double in wild-type cells. This is similar to what has been observed in sulfolipid-deficient mutants in other organisms such as Arabidopsis, which showed impaired growth after severe phosphate limitation (Y u et al., 2002), and in the photosynthetic purple bacterium Rhodobacter sphaeroides (Benning et al., 1993). In addition, Asqdl showed sensitivity to a photosystem II inhibitor under normal growth conditions (Riekhof et al., 2003). This is consistent with another Chlamydomonas SQDG-deficient mutant, hf-Z, which was first discovered as a high chlorophyll fluorescence mutant, and was later found to be impaired in photosystem II stability and showed increased sensitivity to a PSI] inhibitor—which could be partially restored by SQDG addition (Minoda et al., 2002, Minoda et al., 2003, Sato et al., 19953, Sato et al., 1995b). However, whether the hf-2 mutant is impaired in grth during phosphate limitation has not been reported, nor has the exact molecular defect in this mutant been determined. Interestingly, detailed biochemical analysis of the Asqdl mutant also lead to the discovery of the novel sulfolipid derivative, 2’-O-acyl- sulfoquinovosyldiacylglycerol (ASQD), which was also not produced in Asqdl (Riekhof et al., 2003). Due to the loss of both sulfolipids in Asqu, the specific roles played by SQDG and ASQD in Chlarnydomonas and phenotypes associated with Asqdl can only be 124 fully interpreted after the identification and characterization of the acyltransferase catalyzing ASQD production has been undertaken. Phosphatidyl glycerol (PG) is presumably the only major phospholipid component of seed plant thylakoid membranes, and biochemical analysis of thylakoid lipid composition has confirmed this to be the case in Chlarnydomonas (Janero and Bannett, I981b, Mendiola-Morgenthaler et al., 1985). While the gene encoding the final enzyme in PG biosynthesis, phosphatidylglycerolphosphate (PGP) phosphatase, remains unknown in plants and algae (Beisson et al., 2003), the putative genes encoding the enzymes which catalyze the formation of the two intermediates, CDP-DAG synthetase and phosphatidylglycerolphosphate synthase, have been identified (Riekhof et al., 2005b, Riekhof and Benning, 2008), but not yet confirmed. While neither the single gene encoding the CDP-DAG synthetase (CDSI) or the two putative plastid paralogs encoding phosphatidylglycerolphosphate synthase (PGPI/Z) have been studied at the molecular/genetic level, PG deficient mutants, mf I and mf 2, have been isolated and studied in great biochemical detail (Dubertret et al., 1994, Dubertret et al., 2002, Gamier et al., 1987, Gamier et al., 1990, Maanni et al., 1998, Maroc et al., 1987, Pineau et al., 2004). The mf 1, 2 mutants were first isolated as low fluorescent strains lacking fimctional PSII as well as an oligomeric form of the light-harvesting chlorophyll antenna (CPII) (Dubertret et al., 1994, Maroc et al., 1987). It was also shown that both mfI and mf 2 contained ~30 % of wild-type PG levels and lacked A3-trans-hexadecenoic acid (16: 11‘3”” [carbons : double bonds Apos"'i°"""’]) (Dubertret et al., 1994, Maroc et al., 1987), a fatty acid that is specifically esterified to chloroplastic PG in both Arabidopsis and Chlarnydomonas (Browse et al., 1985, Gamier et al. , 1987, Giroud et al., 1988). Addition 125 A3trans of a preparation of spinach leaf PG containing 16:1 to mf-2 cells restored the ability to form oligomeric CP 11, while 18:0 PG additions did not, and 16:0 PG did only weakly so (Dubertret et al., 1994, Gamier et al., 1990). A PG-deficient mutant in Arabidopsis, pgpl , which is defective in the chloroplastid isoform of PGP synthase has been found to be photosynthetically impaired with decreased quantum yield through PSII, but did not lack 16:1A3'm’" PG (Babiychuk et al., 2003, Hagio et al., 2002, Xu et al., 2002). Similarly, two Synechocystis PG deficient mutants showed altered PSII activity and required exogenous PG addition for phototropic growth (Hagio et al., 2000, Sato et al., 2000). Taken together with the contrasting findings from analyses of the Arabidopsisfad4 mutant, which lacks 16:1A3’m'” fatty acid, but is otherwise not affected in chloroplast PG content and also shows no apparent photosynthetic defects, it can currently only be concluded that in general PG plays an important role in photosynthetic membrane biogenesis and function, and it seems possible that the 16:1A3'm'” PG form could be essential in some organisms (e.g. Chlarnydomonas), but is of conditional importance or dispensable in others. It is certain however, that the elucidation of the exact molecular defects in the Chlamydomonas mfl, 2 mutants, and the identification of the genes encoding FAD4 activity as well as the elusive plant/algal PGP phosphatase which catalyzes the final step in PG biosynthesis, will be prerequisite to gaining a better understanding of the roles PG plays in the photosynthetic membranes in various species. Toward this end, the gene affected in the Arabidopsisfad4 16:1A'm’” deficient mutant was . cloned (Gao et al., 2009), and appears to belong to a distinct class of desaturases, as described below. 126 Extrachloroplastic membrane lipid metabolism. In eukaryotes the bulk of extraplastidic membrane glycerolipids are assembled in the ER from acyl-CoA thioesters which are formed from free fatty acids after their liberation from acyl-ACPS in the plastid (see Figure 4.1). While other extraplastidic sites for lipid synthesis are known (e. g. mitochondria), the ER-localized pathway is predominant, and in most plants the ER lipid assembly pathway significantly contributes to thylakoid membrane biogenesis. As such, a discussion of the analogous pathways in Chlarnydomonas is merited. As mentioned above, C hlamydomonas lacks the capability for PC biosynthesis (Giroud et al., 1988) and genes predicted to encode enzymes involved in PC biosynthesis are not present in its genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). Instead, it contains the non- phosphorous zwitterionic betaine lipid DGTS in its membranes (Eichenberger and Boschetti, 1977, Janero and Barrnett, 1982) which has similar structural (see Figure 2), and hence biophysical properties to PC (Sato and Murata, I991). DGTS has also been found in other algal species, (Eichenberger, 1982), prokaryotes like the purple bacterium Rhodobacter sphaeroides, (Benning et al., 1995, Hofrnann and Eichenberger, 1996), and in non-seed plants such as ferns, (Eichenberger, 1993, Sato and Furuya, 1983), but appears to be absent in seed plants. Labeling studies suggest that the biosynthesis of DGTS is similar in all organisms studied (Hofmann and Eichenberger, 1996, Sato, 1988, Sato and Kato, 1988, Vogel and Eichenberger, 1992). It begins with the transfer of the 3- amino 3-carboxypropyl residue from S-adenosylrnethionine (AdoMet) to DAG catalyzed by AdoMetzDAG 3-amino-3-carboxypropyltransferase activity followed by successive methylation of the amino group by an AdoMet-dependent N-methyltransferase . The two genes encoding these catalytic 127 O \ O\P/ i + \ / \\/\O/ O/\l/\O R1 0 R2 PC \11/ O 000' o \+NJ\/\O/\‘/\O/lkR1 0 R2 DGTS T 0 Figure 4.2. Structures of PC and DGTS. A comparison of DGTS and PC structure, where sn-I and sn-2 acyl chains are represented by R. and R2, respectively. 128 activities, BtaA and BtaB, were first identified in R. sphaeroides (Klug and Bennin g, 2001, Riekhof et al., 2005a). More recently, a single gene sufficient for DGTS biosynthesis in Chlamydomonas, Btal, was identified in the genome (Riekhof et al., 2005b). The encoded protein Btal is similar in its N-terminal domain to bacterial BtaB and in its C-terminal domain to BtaA and the predicted catalytic function of each Btal domain was confirmed by mutagenesis (Riekhof et al., 2005b). The bifunctionality observed in Btal as a fusion of two prokaryotic enzyme activities functional in the same pathway into a single polypeptide is a common theme in plants (Moore, 2004), and could represent an improvement in DGTS biosynthesis by eliminating the need for coordinated regulation of two independent gene products or by permitting substrate channeling. In addition, the presence of DGTS and concomitant lack of PC is perhaps related to the absence of additional galactolipid biosynthetic pathways seen in seed plants, e.g., MGD2, 3 and DGD2 in Arabidopsis (Benning and Ohta, 2005). As this alternative galactolipid pathway is believed to be involved in replacing phospholipids in extraplastidic membranes with DGDG during phosphate deprivation, the constitutive replacement of PC with the non-phosphorous DGTS has perhaps obviated the need to replace PC by extraplastidic DGDG. Interestingly, DGTS was tentatively identified as a component of purified Chlarnydomonas thylakoids (Janero and Barrnett, 1981b, J anero and Barrnett, 1982) and chloroplast envelope membranes (Mendiola-Morgenthaler et al., 1985). However, whether DGTS is indeed present in the chloroplast membranes of Chlamydomonas or plays a specific function in thylakoids remains to be confirmed. Phosphatidylserine (PS) is a minor component of extraplastidic membranes of plants and can be decarboxylated to phosphatidylethanolamine (PE) in plants and other 129 organisms (Nerlich et al., 2007, Vance, 1990). However, PS is absent in C hlamydomonas membranes (Giroud et al., 1988) and genes encoding the phosphatidylserine synthase or relevant phospholipid base exchange enzymes were not detected in the genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). AS such, the biosynthesis of PE, which is a known component of extraplastidic membranes in C hlamydomonas, is likely only carried out by a single pathway (Figure 4.1). Genes encoding a serine decarboxylase which produces ethanolamine, an ethanolamine kinase and CTP : phosphoethanolamine cytidylyltransferase—the combined activities of which produce CDP-ethanolamine, and lastly, a CDP-ethanolamine : DAG ethanolamine phosphotransferase which produces PE have been identified in the genome (Riekhof et al., 2005b, Riekhof and Bennin g, 2008). Recently, the gene encoding the CTP : phosphoethanolamine cytidylyltransferase has been characterized by heterologous expression in Escherichia coli, and the expression of the respective gene was found to be up-regulated during the reflagellation of Chlamydomonas cells (Yang et al., 2004). Phosphatidylinositol (PI) is a minor component of Chlamydomonas membranes, and genes required for its biosynthesis are present in the genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). The PI biosynthetic enzymes of Chlamydomonas have been studied at the biochemical level, and PI synthesis was found to be highest in the microsomal fraction, suggesting its association with the ER (Blouin et al., 2003). Extraplastidic PG biosynthesis is known to be associated with both the ER and mitochondria, and three isoforms of phosphatidylglycerolphosphate synthase are encoded in the genome, each with differential targeting prediction probabilities for subcellular localization to the mitochondria, chloroplast or cytosol (Riekhof et al., 2005b, Riekhof 130 and Benning, 2008). However, a detailed analysis of these proteins and their respective genes is not yet available in C hlamydomonas. Fatty Acid Desaturation. Biochemical studies in Chlamydomonas have indicated that further desaturation of 16:0 and 18:1 acyl groups occurs after the production of the major glycerolipids, in a manner similar to plants (Giroud et al., 1988). Furthermore, the Chlarnydomonas fatty acid profile is known to change markedly in response to different environmental conditions including C02 concentration as well as nitrogen and phosphorous limitation (Tsuzuki et al., 1990, Weers and Gulati, 1997). The elucidation of fatty acid desaturase (FAD) genes in Arabidopsis is a classic example of the power of genetic and molecular biological approaches in solving biological problems which prove to be largely intractable through a strictly biochemical approach (Browse et al. , 1985, Wallis and Browse, 2002). The identification of many of the Chlamydomonas desaturase gene candidates by their similarity to Arabidopsis orthologs, combined with a handful of studies of C hlamydomonas desaturase mutants and characterization of cloned FAD genes, has provided a reasonable picture of the fatty acid desaturation pathways in this alga (Figure 4.3). Putative orthologs for the plastidic desaturases encoded by Arabidopsis FAD5 (MGDG palmitate-A7-desaturase), FAD6 (m6-desaturase), and FAD? or FAD8 (encoding m3-desaturase isozymes) are present in the C hlamydomonas genome (Riekhof et al. , 2005b, Riekhof and Benning, 2008). Of these, only the Chlarnydomonas FAD6 gene has been studied to date at the molecular/genetic level (Sato et al., 1995b, Sato et al., 1997). 131 Figure 4.3. Overview of acyl-chain desaturation in Chlarnydomonas. Glycerolipid abbreviations are the same as those in Figure 4.]. Fatty acids are referred to by the standard abbreviation “carbon atomszdouble bonds.” Specifically, common names of these fatty acids include 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 16:1, palmitoleic acid; 16:1”, A3-trans hexadecenoic acid; 18:2, linoleic acid; (118:3, linolenic acid (18:3A9'12’”); i18:3, pinolenic acid (18:3A5‘9’”); 18:4, coniferonic acid. The desaturases that catalyze the enzymatic steps in the plastid (24-29) and the ER (30-32) are indicated by boxed letters: 24, stearoyl-ACP desatruase (des.); 25, PG-specific-16:O- A3trans des.; 26, MGDG-specific-l6:0-A7 des.; 27, plastidic (0'6 des.; 28, plastidic (0-3 des.; 29, MGDG-16C-A4 des.; ER 03-6 des.; 31, A5 des.; 32, ER (0-3 des. 132 l I l i l Lipid Bros nthesls PG SQDG MGDG DGDG (16:0)18:1/16:O 18:1/16:0 18:1/16:0 18:1/16:0 ranger“ 1:33am 133237311115;7| refiso 18:2/16:1” 018:3/1620 18:2/16:2 0181311610 018:3/16z1A3‘ mas/16:3 lk Plastid 018:3116z4 J l \\‘_ _ -_ _ .-_..-"/ ER DGTS PE 16:0/18:1 18:0/18:1 16:0/18:2 182011822 fl 18:0-ACPE—b1821-ACP momma \a 16:0/i18:3 16:0/018:3 .VQ 18:0/18:4 133 \IE 18:0/118:3 18:0/a IE 18:0/18:4 18:3 El The hf-9 mutant was first isolated by its high chlorophyll fluorescence phenotype, and detailed lipid analysis revealed an apparent defect in m6-desaturase activity as it showed marked decreases in both 16- and l8-carbon polyunsaturated fatty acyl groups, with concomitant increases in 16:1A7 and 18:1 A9 (Sato et al., 1995b). The hf-9 mutant had an increased doubling time and showed reduced photosynthetic 02 evolution as well as an altered chloroplast ultrastructure (Sato et al., 1995b). The C hlamydomonas FAD6 gene (first described as DES6) was subsequently cloned and found to be highly similar to cyanobacterial A12- and seed plant (rm-desaturases, and was also Shown to complement the hf-9 mutant desaturation defects (Sato er al., 1997). However, it did not restore the photosynthetic defects, suggesting that these phenotypes arose from a mutation in another gene (or possibly from multiple loci). As such, the roles of polyunsaturated fatty acids (PUFAS) in the assemblage or maintenance of optimally functioning photosynthetic membranes in Chlarnydomonas cannot be easily deduced from analysis of hf-9. The function of PUFAS in this regard have been studied and found to differ in mutants of both plants and cyanobacteria. The Arabidopsisfad6fad2 double mutant, which lacks both plastidic and ER (rm-desaturases exhibited severe grth and photosynthetic defects (McConn and Browse, 1998). In contrast, a mutant of Synechocystis Sp. PC6803 lacking PUFAS had no observable photosynthetic defects under normal growth conditions (Gombos et al., 1992). Thus, it still remains to be determined whether the importance of PUFAS in photosynthetic membrane function in C hlamydomonas is more similar to that of seed plants or cyanobacteria. As noted above, a candidate desaturase producing 16:1A3’m'" specifically on plastidic PG, was identified (Gao et a1. , 2009) by map-based cloning of the genetic lesion in the Arabidopsisfad4 mutant (Browse et al., 1985). The 134 FAD4 protein is distinct from all other known acyl-chain desaturases with respect to position and number of proposed catalytic residues, which may be related to the fact that this enzyme produces a trans rather than cis desaturation product. A candidate FAD4 ortholog has been identified in the most recent draft of the Chlarnydomonas genome (v 4.0, http://genome.jgi-psf.org/chlamy/chlamyhome.html), but has not yet been characterized (S. Zaiiner and C. Benning, pers. comm.) However, the gene list of chloroplast desaturases in C hlamydomonas remains incomplete, as a gene encoding A4 desaturase activity, which is specific for MGDG-esterified l6-carbon acyl groups based on biochemical studies (Giroud et al., 1988), and presumably produces both the 16:3’34’7‘lo and 16:4‘54’7"°'l3 in Chlarnydomonas is currently unidentified. The extraplastidic 036- and co3-desaturases which produce 182“”12 and 18:3 ”'2’” are encoded by FAD2 and FAD3, respectively in Arabidopsis, and putative orthologs of these genes have been identified in the Chlamydomonas genome (Riekhof et al., 2005b, Riekhof and Benning, 2008). The extraplastidic Chlarnydomonas lipids A5,9,12 3 DGTS and PE have been shown to contain significant amounts of 18: and 1824559”15 esterified to the respective sn-2 positions of the glycerol back bone (Giroud et al. , 1988); these A5-unsaturated fatty acids are also found in gymnosperms (Mongrand et al., 2001, Wolff and Christie, 2002). Recently a Chlarnydomonas gene, CrDES, encoding a “fi'ont-end” type AS-desaturase was identified by a bioinformatics approach through its similarity to a known A5-desaturase from the liverwort Marchantia polymorpha (Kajikawa et al., 2006). Heterologous expression of CrDES in Pichia pastoris and analysis of desaturase activity indicated while the primary substrates were 18:21‘9’12 and 18:3 ”’12”, low but detectable levels of endogenous 18:1” desaturation 135 were also observed (Kajikawa et al., 2006). Transgenic tobacco plants constitutively expressing the C rDES gene exhibited strikingly high levels of 18:3’35‘9‘12 and 18:4’35‘9‘n‘l5 (which are normally absent), with the highest combined yield reaching ~45% of leaf total fatty acids, and no apparent morphological phenotypes (Kajikawa et al., 2006). The biological roles of these A5-unsaturated fatty acids in the organisms which produce them are largely unknown, and the identification of the CrDES gene responsible for 1823“"'2 and 18:4’35‘9’12’ls production in Chlarnydomonas will allow for this gene to be targeted for suppression through RN Ai technology. Differences in ER-to-chloroplast lipid trafficking in plants and Chlarnydomonas The diacylglyceryl backbone moieties that feed into thylakoid lipid biosynthesis in many plants are derived from two pathways (Mongrand et al., 1998), termed the plastid and the ER pathways. This hypothesis was first formulated by Roughan and coworkers based on labeling experiments (Roughan et al. , 1980, Roughan and Slack, 1982) and later confirmed by analysis of Arabidopsis mutants (Browse and Somerville, 1991, Wallis and Browse, 2002). The relative contribution of each pathway is evident in the fatty acid composition of chloroplast-specific lipids (e. g. MGDG), where the ratio of 16:3 (plastidic pathway) to 18:3 (ER pathway) at the sn-2 position is diagnostic (Heinz and Roughan, 1983), and fluxes through the two pathways have been determined (Browse et al., 1986). Many plant Species have lost the plastidic pathway for providing DAG moieties for biosynthesis of the dominant galactoglycerolipids, but nearly all plant species reported derive at least a fraction of their thylakoid lipids from precursors assembled at the ER (Mongrand et al., 1998) requiring lipid precursor import into the plastid. Recent studies have identified genes encoding a putative ABC-type transporter 136 complex localized to the chloroplast inner envelope, which is required for ER-to-plastid lipid trafficking in Arabidopsis, by analysis of mutants in the TGD1,2,3 genes (Awai et al., 2006, Lu etal., 2007, Xu et al., 2003). In contrast, detailed compositional analysis of lipids and labeling studies suggest that in Chlamydomonas all thylakoid lipids are assembled de novo in the plastid (Giroud et al., 1988). Intriguingly, orthologs of both the Arabidopsis TGDl and 2 proteins are encoded in the Chlarnydomonas genome, suggesting they be essential for lipid trafficking within the plastid in the absence of ER glycerolipid import, or alternatively, they are not functioning in lipid transport in Chlarnydomonas. This could also be related to the lack of PC biosynthesis in Chlamydomonas, and its (presumed) functional replacement with DGTS (Eichenberger and Boschetti, 1977). PC is central to lipid metabolism in developing seeds or leaves where it serves a substrate for fatty acid modifying enzymes such as desaturases (Browse and Somerville, 1991, Ohlrogge and Browse, 1995, Wallis and Browse, 2002) or possibly as the lipid transferred between the ER and the plastid (Benning, 2008, Jouhet et al., 2007). Thus, it is possible that the lack of PC and the lack of trafficking of lipid precursors from the ER to the plastid in Chlarnydomonas are related if PC is a critical intermediate in ER-to-plastid lipid trafficking. Because in the betaine lipid, DGTS, the head group moiety is ether-linked to the diacylglyceryl moiety, Chlarnydomonas might lack an enzyme to break this ether linkage. This ether linkage is more stable than the phosphate ester linkage in phosphoglycerolipids. Therefore, the conversion of DGTS into the galactoglycerolipid precursor diacylglycerol might not be possible in C hlamydomonas. 137 Neutral glycerolipid metabolism. Many algae accumulate significant quantities of neutral lipids (predominately TAG) under certain conditions, and thus have long been considered an attractive source for bio-based fuels. While there has been a resurgence in interest in algal biofuel production (Hu et al., 2008), this concept was intensely studied under the two decade spanning Dept. of Energy funded Aquatic Species Program (ASP) that initiated in the late 1970’s (Sheehan et al., 1998). Through screening thousands of algal strains, many species were identified that could accumulate TAG up to 75 % dry weight (Ben-Amotz et al., 1989, Bigogno et al., 2002, Chisti, 2007). In most algae species, TAG accumulates in response to environmental stress, suggesting that triacylglycerol plays a role in microalgae beyond energy storage (Chisti, 2007, Hu et al., 2008, Shanklin and Somerville, 1991). Most studies focused on nutrient deprivation, with an emphasis on nitrogen limitation, where it was found that the chlorophytes Neochloris oleoabundans (Tomabene et al., 1983) and Parietochloris incise (Merzlyak et al., 2007) accumulate TAGS. The eustigmatophyte Nannochloropsis gave similar results (Suen et al., 1987), suggesting that this response to N-limitation may be a common phenomenon in algae—given the large evolutionary divergence between the Chlorophycae and Bustigrnatophycae (Reyes-Prieto et al., 2007). Other nutrient deficiencies are also known to cause TAG accumulation. For example, silicon limitation results in increased TAG content in the diatom Cyclotella cryptica (Roessler, 1988). Light intensity and temperature are also known to affect the accumulation of TAG in many algal Species. Changes in temperature result in species specific-alterations in lipid accumulation, with some algae showing increased TAG content, and others showing decreases (de Swaaf et al., 1999, Dempster and Sommerfeld, 1998, Richardson et al., 138 1983). Growth of many algal species under high light intensity results in increased TAG to total lipid ratios (Khotimchenko and Yakovleva, 2004, Khotimchenko and Yakovleva, 2005, Zhekisheva et al., 2002). The chlorophyte Dunaliella bardawil accumulates TAG under high light Stress along with B-carotene, which was proposed to protect chloroplasts from photo-oxidative damage (Ben-Amotz et al. , 1989, Rabbani et al., 1998). While Chlamydomonas reinhardtii was largely considered a poor lipid accumulator, this species can accumulate significant amounts of TAG during nitrogen or phosphorous deprivation (W eers and Gulati, 1997). Despite these efforts, little is currently known about algal TAG biosynthesis at the molecular level, including that in the genetic model alga Chlarnydomonas. Given the above-described Similarities between Chlamydomonas and plant membrane glycerolipid metabolism, it seems likely that many general aspects of TAG metabolism are also shared. A major pathway for TAG biosynthesis in many organisms, including plants, involves the step-wise acylation of a glycerol backbone (i.e. the Kennedy pathway; see Figure 1) (Kennedy, 1961). However, several studies in many different plant species have also indicated that DAG derived from the PC pool also contributes substantially to TAG biosynthesis (Bates et al., 2009, Ohlrogge and Browse, 1995). Clearly, PC plays no role in TAG biosynthesis in C hlamydomonas, and its role in plants is not universal, as studies of avocado mesocarp microsomes indicated that only the Kennedy pathway was active (Stobart and Stymne, 1985). It may be possible that the assumed functional analog of PC in Chlarnydomonas, DGTS, is an intermediate in the biosynthesis of TAG. However, no biochemical studies to determine this have been undertaken to date, and utilization of the DGTS pool to provide DAG precursors would require an as yet 139 unidentified enzyme to remove the ether linked trimethylhomoserine head group. Regardless of the precursors used in forming DAG, the final step is catalyzed by diacylglycerol acyltransferases, or DGATS, which transfer a fatty acid from acyl-CoA to diacylglycerol. DGATs have been isolated and characterized from several plant species, including Arabidopsis (Hobbs et al., 1999, Routaboul et al., 1999, Zou et al., 1999), maize (Zheng et al., 2008) and castor beans (Kroon et al., 2006). The Chlarnydomonas genome contains a number of putative DGAT isoforms, which all appear to be of the type 11 class of DGAT enzymes (Lardizabal et al., 2001), and remain to be studied in molecular detail (Riekhof and Benning, 2008; Miller and Benning, unpublished). Chlarnydomonas may also use an alternate pathway for TAG synthesis involving the phospholipid : diacyl glycerol acyltransferases, or PDATS, to generate TAG using a phospholipid as a fatty acid donor, rather than acyl-CoA (Dahlqvist et al., 2000). PDATs have also been found in plants (Stahl et al., 2004), and a putative ortholog was also found in Chlamydomonas (Riekhof and Benning, 2008). Given the induction of TAG biosynthesis by different stresses, it is likely that the mechanism for the regulation of TAG synthesis differs in Chlamydomonas fiom that in seed plants, which often produce oil during a specific phase of their life-cycle and in specialized tissues. Intriguingly, a recent study of chloroplast lipid remodeling during N-deprivation in Arabidopsis revealed the accumulation of TAG and phytyl esters in the chloroplast (Gaude et al. , 2007), suggesting that TAG accumulation may be a common response in photosynthetic organisms to carbon-nitrogen imbalances brought about by N-limitation. 140 Perspectives Interest in microalgal lipid metabolism is experiencing a renaissance due to the increasing need for replacing fossil fuels with bio-based renewable resources. This, coupled with the ever-decreasing cost of DNA sequencing technologies, will undoubtedly result in the sequencing of a number of algal genomes. Thus, comparative genomics may become a powerful tool in elucidating factors required for TAG biosynthesis in algae. Moreover, such studies in other algal Species may reveal distinct mechanisms of lipid metabolism that have evolved separately in this highly divergent group of organisms. The current availability of the C hlamydomonas genomic sequence (Merchant et al., 2007), and its corresponding molecular toolkit has indeed made the application of knowledge on well studied lipid metabolism in seed plants to this model alga relatively facile using comparative genomics. As such, the potential for applying insight gained from studying lipid metabolism in Chlarnydomonas in gene discovery and the rational engineering of other algal species to produce biofuels is promising. References Awai, K., Marechal, E., Block, M. A., Brun, D., Masuda, T., Shimada, H., Takamiya, K., Ohta, H., and J oyard, J. (2001) 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. U. S. A 98, 10960-10965. Awai, K., Xu, C., Tamot, B., and Benning, C. 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(2008) A phenylalanine in DGAT is a key determinant of oil content and composition in maize. Nat. Genet. 40, 367-372. Zheng, Z., Xia, Q., Dauk, M., Shen, W., Selvaraj, G., and Zou, J. (2003) Arabidopsis AtGPATl , a member of the membrane-bound glycerol-3-phosphate acyltransferase gene family, is essential for tapetum differentiation and male fertility. Plant Cell 15, 1872- 1887. Zou, J., Wei, Y., Jako, C., Kumar, A., Selvaraj, G., and Taylor, D. C. (1999) The Arabidopsis thaliana TAGl mutant has a mutation in a diacyl glycerol acyltransferase gene. PlantJ. 19, 645-653. 153 Chapter 5 Analysis of triacylglycerol accumulation in the green alga Chlamydomonas reinhardtii during nitrogen deprivation: Identification and characterization of an algal specific lipid droplet protein MLDPl ' This work was published in: Moellering, E. R. and C. Benning. (2010) RNA interference Silencing of a major lipid droplet protein affects lipid droplet size in C hlamydomonas reinhardtii. Eukaryot. Cell 9:97-106. 154 Abstract Eukaryotic cells store oils in the chemical form of triacylglycerols in distinct organelles, ofien called lipid droplets. These dynamic storage compartments have been intensely studied in the context of human health and also in plants as a source of vegetable oils for human consumption and for chemical or biofuel feedstocks. Many microalgae accumulate oils, particularly under conditions limiting to growth and, thus, have gained renewed attention as a potentially sustainable feedstock for biofuel production. However, little is currently known at the cellular or molecular levels with regard to oil accumulation in microalgae, and the structural proteins and enzymes involved in the biogenesis, maintenance, and degradation of algal oil storage compartments are not well studied. Focusing on the model green alga Chlamydomonas reinhardtii, the accumulation of triacylglycerols and the formation of lipid dr0plets during nitrogen deprivation are investigated Mass spectrometry identified 259 proteins in a lipid droplet enriched fraction, among them a major protein, tentatively designated Major Lipid Droplet Protein (MLDP). This protein is specific to the green algal lineage of photosynthetic organisms. Repression of MLDP gene expression using an RNAi approach led to increased lipid droplet size, but no change in triacylglycerol content or metabolism was observed. 155 Introduction Triacylglycerols (TAGS) are stored in lipid droplets which are subcellular structures in specialized cells ubiquitous to eukaryotes, but have more recently also been identified in some prokaryotes (Murphy, 2001). In plants and animals, lipid droplets are surrounded by cytosol and are believed to bud off the endoplasmic reticulum (Huang, 1992, Murphy, 2001). While traditionally considered merely as storage compartments, recent studies suggest that lipid droplets in animals play important additional roles in lipid homeostasis, and protein storage (Cermelli et al., 2006). In oilseed plants, TAG accumulated in seeds is used as a reservoir of energy and membrane lipid building blocks to support rapid growth after germination (Huang, 1992). Many green algae are capable of accumulating large amounts of TAG in lipid droplets, particularly as a result of abiotic stresses such as nutrient deprivation or high-1i ght exposure. Although TAG metabolism in algae has not yet been extensively studied at the biochemical or molecular level, it is proposed that TAG turnover contributes primarily to the assembly of membrane lipids to facilitate rapid cell division after the cessation of nutrient limitation (Hu et al., 2008, Thompson, Jr., 1996). The general structure of lipid droplets is conserved in different species with a globular neutral lipid core enclosed by a membrane lipid monolayer (Murphy, 2001). In addition, Specific proteins are associated with lipid droplets and play important roles in lipid droplet structure and function. A number of recent proteomic studies of lipid droplets from different animals and tissues (Cermelli et al., 2006, Walther and Farese, Jr., 2009), yeast (Athenstaedt et al., 1999), and from plants (Jolivet et al., 2004, Katavic et al., 2006) have revealed that the lipid droplet-associated proteins of these organisms are 156 quite distinct. For example, the abundant lipid droplet proteins in animals—the so called “PAT” family of proteins comprised of perilipin, adipose differentiation-related protein (ADRP) and T1P47 (Londos et al., 2005), have no apparent orthologs in the desiccating seed plant Arabidopsis thaliana; conversely, the oleosins which coat the oil bodies of Arabidopsis and many other seed plants are not found in animals (Murphy, 2001). Reverse genetic Studies of these proteins have helped to elucidate the role of A. thaliana oleosins in regulating lipid droplet size and preventing droplet fusion (Shimada et al. , 2008, Siloto et al., 2006), or that of mouse adipocyte perilipin in regulating lipolytic activity at the lipid droplet surface (Tansey et al., 2001). Moreover, recent genome-wide RNAi screens in Drosophila cells implicated 1.5-3.0 % of all genes as directly or indirectly involved in lipid droplet formation and/or regulation, and resulted in the identification of a new role for Arfl-COPI vesicular transport machinery in regulating droplet morphology and lipid utilization (Beller et al., 2008, Guo et al., 2008). In contrast, few molecular details are known about algal lipid droplet biogenesis although many TAG-rich algal species have been described (Hu et al., 2008). Our efforts to identify proteins related to the PAT protein family or oleosins in the Chlamydomonas reinhardtii genome (Merchant et al., 2007) or genomes of other green algal and diatom species including Thalassiosira pseudonana, Volvox carteri, and Chlorella sp.NC64A revealed no putative algal orthologs. In order to identify both potentially novel and conserved proteins which function in algal lipid droplet biogenesis, we studied the accumulation of TAG in lipid droplets of nitrogen-(N)-lirnited C. reinhardtii cells, and identified candidate lipid-droplet associated proteins by mass spectrometry. 157 Materials and Methods Strains and growth conditions. Chlamydomonas reinhardtii strains dw15 (cw15, nit], mt+) and CC-125 (nitl, nit2, mt+) were grown in Tris-acetate-phosphate (TAP) medium with N114Cl at 10 mM, in liquid cultures or on agar-solidified plates under continuous light (70—80 umol m"2 s") as previously described (Riekhof et al., 2005). Strain CC-125 was used in generating the data shown in Figure 5.1B and Figure 5.2, and de5 or transgenic derivatives thereof were used in the remainder of this study. To induce N- lirnitation, mid-log phase cells grown in 10 mM NH4C1 TAP were centrifuged for 5 minutes at 5,000 x g, washed once in TAP lacking NH4C1 or any other N-source (TAP- N), and resuspended in TAP-N. Cells were counted with a hematocytometer. Isolation of lipid droplets from N-limited C. reinhardtii. Cells grown in 500 ml TAP-N medium for 24 h were centrifuged 3,000 x g for 5 min and resuspended in 10 ml of isolation buffer (50 mM HEPES, pH 7.5, 5 mM MgC12, 5 mM KCl, 0.5 M sucrose, 1 mM phenylmethylsulfonylfluoride, 1 mM 2,2’-dipyridyl, with 1x protease inhibitor cocktail [Rocbe, Branchburg NJ ]). Cells were disrupted with the aid of a Potter homogenizer, using 20 strokes as well as sonication at the lowest energy setting (3 W output, Misonex Sonicator 3000, Newton, CT) for 5 sec. The homogenate was overlayed with buffer 2 (same as isolation buffer above, but lacking 0.5 M sucrose) and centrifuged at 100,000 x g for 30 min. Lipid droplets floating on the overlay buffer were removed with a bent Pasteur pipette and diluted lO-fold with washing buffer (same as isolation buffer, but with KCl increased to 150 mM). The suspension was overlayed with buffer 2 and centrifuged as described above. This washing process was repeated three times, and lipid 158 droplets were suspended in buffer 2 and stored at -80 °C. Preliminary attempts to isolate lipid droplets with this method from cells deprived of nitrogen for 2 or 3 days resulted in lipid droplets with significantly increased chlorophyll contamination compared to lipid droplets fiom 24 h -N cells (data not shown)»as such lipid droplets from 24-h -N grown cells were used throughout in this study. Thylakoid membranes were isolated according to (Chua and Bennoun, 197 5), and enriched eyespots were prepared as previously described (Schmidt et al., 2006). Mass spectrometric identification of lipid droplet proteins. Proteins were precipitated as previously described (Schmidt et al., 2006) and solubilized in SDS sample buffer by sonication. Proteins were separated on 4-20% SDS-PAGE gels (Biorad, Hercules, CA) and stained with EZBLUETM gel staining reagent (Sigma-Aldrich, St. Louis, MO). The lane containing lipid droplet proteins was sliced into 15 bands, which were processed for in-gel trypsin digestion as previously described (Shevchenko et al., 1996) and analyzed by LC-MS/MS by the Michigan State University Research and Technology Facility. The extracted peptides were re-suspended in a solution of 2% acetonitrile/O.l% trifluoroacetic Acid to 20 pl. From this 10 pl were automatically injected by a Waters nanoAcquity Sample Manager (Waters, Milford, MA) and loaded for 5 minutes onto a Waters Symmetry C18 peptide trap (Sum, l80um x 20mm) at 4 uI/min in 2% aceto nitrile/O. 1% formic acid. The bound peptides were then eluted onto a Waters BEH C18 nanoAcquity column (1.7 pm, lOOum x 100mm) and eluted over 35 minutes with a gradient of 2% B to 35% B in 21min, 90% B from 21- 24 minutes and 5% B after 24.1 minutes at a flow rate of 0.6 111/min. Separations were done using a Waters nanoAcquity UPLC (Buffer A 159 = 99.9% Water/O. 1% F ormic Acid, Buffer B = 99.9% Acetonittile/O. 1% F onnic Acid) and sprayed into a Thermo Fisher Scientific (Waltham, MA) LTQ mass spectrometer using a Michrom ADVANCE nanospray source. Survey scans were taken in the ion trap and the top five ions from each survey scan were automatically selected for high resolution Zoom scans followed by low energy collision induced dissociation (CID). The resulting MS/MS Spectra were converted to peak lists using BioWorkS Browser v 3.3.1 (Thermo Fisher Scientific) using the default parameters and searched against the Chlamydomonas reinhardtii database, v3.0, downloaded from The DOE Joint Genome Institute (www.jgi.doe.gov), using the Mascot searching algorithm, v 2.1 (www.matrixscience.com). The Mascot output was then analyzed using Scaffold (wwwproteomesoftwarecom) to probabilistically validate protein identifications using the ProteinProphet computer algorithm (Nesvizhskii et al., 2003). Assignments validated above the Scaffold 95% confidence filter were considered true. Mascot parameters for_ database searching allowed up to 2 missed tryptic sites, variable modification of oxidation of methionine and carbarnidomethyl cysteine, a peptide tolerance of +/- 100 ppm, a MS/MS tolerance of 0.8 Da, and a peptide charge state limited to +2/+3. Fatty acid and lipid analysis. Various cellular and sub-cellular fractions were extracted into methanol:chloroform:formic acid (2: 1 :0.1 vol) followed by phase separation by the addition of l M KC1, 0.2 M H3PO4. The organic phase was spotted on ammonium sulfate impregnated silica Si250 plates, and TAG was separated with petroleum etherzdiethyl etherzacetic acid (80:20:1 vol). After visualization by brief iodine staining, TAG bands were quantitatively scraped from the plates and processed for fatty acid methyl 160 esterification (FAME) and quantified by gas chromatography as previously described (Rossak et al., 1997). For total cellular fatty acids, the total cellular extract from a known number of cells was processed directly for FAME as described above. Construction of vectors and transformation of C. reinhardtii. All PCR reactions were performed using the Phusion® High-Fidelity DNA polymerase (Finnzymes, Wobum, MA) according to the manufacturer’s instructions, all ligation reactions were performed with T4 DNA ligase (Invitrogen, Carlsbad, CA), and the sequences of the modified regions of all final expression vectors were confirmed by DNA sequencing at the Michigan State University Research Technology Support Facility. In order to construct the pNITPROl vector, the C hlamydomonas RBCSZ 3’UTR was amplified from genomic DNA with primers P1 5’-GTTGGTACCTGATAAGGATCCCCGCTC-3’ and P2 5’- GTTGTAAAACGACGGCCAGTGCC-3’ and ligated into Kpn] digested pBluescript II KS+ (Stratagene, Agilent Technologies, Santa Clara, CA) after digestion with the same enzyme; a clone containing the desired orientation of the insert was identified by PCR amplification using P2 5’-GTTGTAAAACGACGGCCAGTGCC-3’ and P3 5’- TAATACGACTCACTATAGGG-3’ (the T7 sequencing primer), and the resulting plasmid was designated pEMl. The C hlamydomonas N] T l promoter region was amplified from the vector pMN24 (Fernandez et al., 1989) using primers P4 5’- CGGT'TCTAGACGTGTATGGCT'I'TGGCTAC-3’ and P5 5’- GGTAACTAGTACTGGCAGGATTCGGCTG-3’. This amplicon was cut with Pcil and Spel and ligated into pRBCSZ after its digestion with the same enzymes; the resulting plasmid was designated pEM2. PCR primers P6 5’- 1'1 IGCTCACATGTACAGGTG-3’ 161 and P7 5’-CCCCCATATGACCCGCTTCAAATACGC-3’ were used to produce a NIT 1 promoter-MCS-RBCS2 3’UTR region of pERM2 amplicon, which was cut with Pcil and ligated into a Pch/Swal digested pSP124S—a plasmid containing the ble selectable marker gene for Chlarnydomonas transformation. To generate an MLDP RNAi construct, primers P8 5’- GCGCACTAGTCTGGAAAGCCTCTGAAGCAC-3’ and P9 5’- CCTTCCATGGGGTCGTAGTACTTGTCGGCG-3’ were used to amplify a sense amplicon from genomic DNA, while P10 5'- CCCCCTGCAGCTGGAAAGCCTCTGAAGCAC-3’ and P11 5’- GGAACCATGGTCGGTGCTCCGGAGAATCTT-3 ’ were used to amplify the antisense amplicon from cDNA. The sense and antisense amplicons were double digested with Spel/Neal and PstI/Nco], respectively, and ligated into Spel/Pstl digested pBluescript II KS+. The formation of the triple ligation product was confirmed by DNA sequencing, and the resulting hairpin insert was excised by Spel/Pstl digestion, and ligated into pNITPROl digested with the same enzymes. The resulting plasmid was designated pMLDP-RNAi. C. reinhardtii strain dw15 was transformed as previously described (Lumbreras et al., 2008), except that vectors were linearized with Pacl, and transformants were selected and maintained on TAP agar plates supplemented with 1 % yeast extract and 10 pg ml"l zeocin. Quantitative Real-time RT-PCR. Total RNA was purified using the RNeasy plant mini kit (Qiagen, Gennantown MA) including the on-column DNAse digestion. 500 ng RNA was reverse transcribed using the RETROscript kit (Ambion, Austin, TX) in a 10 pl 162 reaction. Real-time PCR was performed on an ABI Prism 7000 (Applied Biosystems, Foster City, CA) using the SYBR green PCR master mix with the equivalent input of 50 ng RNA and 0.3 pM each primer (MLDP- F 5’-GGATGCCTGGACCAAGTTC-3’ and MLDP-R 5’-GAGTGCACCAGGAGGTCGT-3’; RACKI-F 5’- GACCACCAACCCCATCATC-3' and RACK I -R 5’-AGACGGTCACGGTGTTGAC- 3’). In all experiments, expression of MLDP was normalized to RACK I (CBLP) expression, a gene commonly used for normalization in C. reinhardtii (Allen et al., 2007). Analysis of RACK] expression by the 2"“ method (Livak and Schmittgen, 2001) resulted in the following fold-changes (average and standard deviation are indicated) in expression when using the average threshold cycle (Ct) value of 10 mM N grown cells: 1.015: 0.17 for 10 mM NI-I4Cl grown cells, and 1.14 :1: 0.1 l, 1.79 :1: 0.72, and 1.62 i 0.28 for cells switched to 0 mM N for l, 2, and 3 days, respectively. The standard curve method for quantification of mRNA abundance was used as previously described (Larionov et al., 2005). Melting curves were performed after PCR to confirm that the amplification product was unique. Fluorescent, confocal, and electron microscopy. For confocal microscopy, cells were re-suspended in phosphate buffered saline (PBS). Cells were stained with Nile Red at a final concentration of 2.5 pg ml'l (from a stock of 50 pg ml’1 in methanol), followed by a 10 minute incubation in the dark. Stained cells were then mixed 1:1 (vol/vol) with 2% low-temperature melting point agarose kept at 38 °C and applied to the microscope slide. Afier the solidification of the agarose, images were captured using a ZeiSS 510 Meta ConforCor3 LSM microscope (Zeiss, Maple Grove, MN). For neutral lipid specific 163 detection of Nile Red fluorescence, the 488 argon laser was used in combination with a 560-615 nm filter; for combined chlorophyll autofluorescence and Nile Red Fluorescence of polar lipids, a 647—753 nm filter was used. Samples were viewed with a 63x oil- irmnersion objective. Postacquisition image handling was done with Zeiss AIM software. For fluorescence microscopy, a Zeiss Axio Irnager M1 microscope was used. Nile Red fluorescence in neutral lipids was detected using a UV light source with a 500 :h 24 urn/542 d: 27 nm exitation/emission filter set. For the measurement of apparent lipid droplet diameter micrographs were analyzed using Image J software (Abramoff et al. , 2004). For electron microscopy, centrifuged cell pellets were processed as previously described (Harris, 1989), and transmission electron micrographs were captured using a JEOLlOO CXII instrument (Japan Electron Optics Laboratories, Tokyo, Japan). Bioinformatics. Sequences were routinely searched using BLAST (Altschul et al., 1997). Sequences were aligned using the Tcoffee multiple sequence alignment program (Poirot et al. , 2003). Hydropathy plots were generated employing the Kyte-Doolittle algorithm (Kyte and Doolittle, 1982) using the ProtScale program at www.cxpasy.ch/tools/protscale.html. The value G in each graph is the grand average of hydropathy value (GRAVY) for each protein and was calculated using the GRAVY calculator program at http://gravylaborfrustdel. 164 Results TAG accumulation following shift to N-limiting medium. While C. reinhardtii does not accumulate TAG under nutrient-replete growth conditions, TAG has been reported to accumulate in this genus after N or phosphorous limitation (Van Donk et al., 1997). In order to optimize conditions for the isolation lipid droplets from C. reinhardtii cells, the time course of N-deprivation and its effect on TAG accumulation and lipid droplet formation was analyzed. The amount of fatty acid esterified in TAG steadily increased over time to approximately 12 %, 45 % and then 65% of total cellular fatty acids 24, 48, and 72 h, respectively, after resuspending the cells in TAP medium lacking a source of N as shown in Figure 5.1A Total fatty acid profiles of N-limited cells showed moderate changes in comparison to N-replete cells, with a slight increase in 16:0 (carbon number: 3A9"2"5 (superscript indicates number of double bonds), and concomitant decrease in 18: position of double bond counting form the carboxyl end); TAG fatty acid profiles of N- replete and N-limited cells were more distinct with a decrease in 16:0 and an increase in 18:1/16:4 in N-lirrrited cells (Table 5.1). Total cellular fatty acids increased on a per cell basis 48 and 72 h after imposing N-limiting conditions (Figure 5.1A), indicating that de novo fatty acid synthesis contributes at least in part to TAG accumulation. A time course of lipid droplet formation in N-deprived C. reinhardtii cells is shown in Figure 5.18. Using the lipid-specific fluorescent dye Nile Red, lipid droplet formation was observed by confocal microscopy adjusting excitation and emission filters to predominately detect fluorescence from the neutral lipid fraction (Elsey et al., 2007). First lipid droplets were observed after 12 h of N-deprivation in distinct locations within 165 > FA (nmol)/ 10 cells to Figure 5.]. Accumulation of triacylglycerols in lipid droplets during N-limitation. (A) Quantification of total cellular fatty acids (black bars) and fatty acids in triacylglycerol (grey bars) of dw15 cells grown in medium with 10 mM NH; (+N), or cells switched (arrow) to medium with no NH! (-N) and grown for 24, 48, or 72 h. (B) Confocal microscopy images of Nile Red-stained cc-125 cells grown in medium with 10 mM NH..‘ (+N) or cells switched (arrow) to medium with 0 mM NH,“ (N) and grown for 12, 24 or 48 h. Differential interference contrast (D), chlorophyll autofluorescence (C), Nile Red fluorescence indicating lipid droplets (L), and all three images merged (M) are shown. The scale bars indicate 5 pm. 166 Table 5.1. Total fatty acid content of whole cells compared to fatty acids bound in triacylglycerols isolated from cells which were grown in medium containing 10 mM NH: (+N), or cells switched to 0 mM N114+ and grown for one day (-N). a Whole cells Whole cells TAG TAG Fattyacid +N -N +N -N 16:0 22.1 1 0.14 26.2 :1 0.70 51.3 1 1.30 37.6 31 3.99 16:1 3.4 1 0.02 2.9 1 0.01 5.4 1 2.37 3.7 :1 0.54 16:2 1.2 1 0.00 1.4 1 0.06 1.6 1 0.96 1.4 1 0.17 16:3 4.5 :1 0.15 3.4 1 0.25 13.9 :1 6.88 3.2 :1 0.27 18:0 2.4 1 0.02 2.5 :1 0.06 0.8 1 0.84 1.4 :1 0.17 18:1,16:4 21.9 1 0.13 21.2 1 0.14 7.9 :1 2.46 22.1 1 8.45 18:2 8.0 3; 0.02 9.9 1 0.06 6.1 :1 3.33 13.6 31 1.51 18:3A5'9‘” 6.7 1 0.01 6.5 1 0.12 4.6 .1 1.62 5.7 1 0.63 18:3A9"2"5 27.9 1 0.10 24.7 1 0.13 7.3 :1 3.69 9.9 31 1.20 18:4 1.9 1 0.02 1.5 :1 0.02 1.1 11 0.16 1.3 :1 0.16 aThe data are presented on a % mol basis where three replicates were averaged and the standard deviation is shown. 167 the cell, and prolonged exposure of the cells to 24 or 48 h of N-deprivation caused an increase in the size and number of lipid droplets. Similarly, transmission electron micrographs revealed several ultra-structural changes that occur over a 72 h time course of N-limitation, including a reduction of stacked thylakoid membranes, accumulation of starch granules, and the appearance of lipid droplets (Figure 5.2). Properties of lipid droplets. Lipid droplets were prepared from 24 h N-deprived cells and compared to isolated thylakoid membranes and a fraction highly enriched in eyespots—a chloroplast localized, carotenoid and lipid-rich organelle (Schmidt et al., 2006). Spectral analyses of the various subcellular fractions indicated the presence of carotenoids (Figure 5.3A). This result was similar to what was previously observed for an enriched eyespot fraction (Schmidt et al., 2006), although when normalized on a per fatty acid basis, carotenoids were much less abundant in lipid droplets. The near absence of chlorophylls in isolated lipid droplets represented by the absorbance maximum at approximately 660 nm visible in the thylakoid fraction (Figure 5.3A) suggested low contamination of the lipid droplets with thylakoid membranes. The ratios of TAG- esterified fatty acids to total fatty acids of lipid droplets and eyespots were approximately 88% and 25%, respectively (Figure 5.3B), corresponding to the higher relative membrane abundance in the eyespots. The TAG fatty acid profile of isolated lipid droplets was very similar to the profile of TAGS isolated from cells deprived of N for 1d, whereas the TAG fatty acid profile of eyespots was quite distinct, being much more enriched for palmitic acid on a mo] percent basis (Table 5.2). Taken together, these results indicated that the lipid droplets were distinct from the lipid globules found in the eyespot fraction, with 168 Figure 5.2. Ultrastructure of cells during N-limitation. Electron micrographs are shown of a representative cc-125 cell grown in medium containing 10 mM NH.+ (0d), and representative cells switched to 0 mM NH4+ and grown for one (1d), two (2d), or three (3d) days respectively. Scale bars represent 2 pm. The Arrows are pointing at: E, eyespot; LD, lipid dr0plets; N, nucleus; P, pyranoid; S, starch granules; T, thylakoid membranes; V, vacuoles. 169 Figure 5.3. Characteristics of isolated lipid droplets. (A) Absorbance spectra of ethanol extracts (96%) normalized on an equal fatty acid basis from thylakoids of cells grown in medium containing 10 mM NH4+ (+N), or cells switched to 0 mM NIH and grown for one day (-N 1d), from enriched eyespot fraction, and from lipid droplets were recorded. (B) Ratios of fatty acids derived from triacylglycerol (TAG FA) to total fatty acids (Total FA) for extracts from whole cells grown in medium with 10 mM N11,.+ (+N), or whole cells switched to and grown in medium with no Nl-I..+ (-N) for 24, 48, or 72 h, the enriched eyespot fraction (Eye), and isolated lipid dr0plets (LD) as indicated. (C) Proteins (30 pg) separated by gradient (4-20%) SDS-PAGE from whole cells (Tot. Prot., total protein), thylakoids, eyespots, and lipid droplets as indicated. The positions molecular weight markers in kDa are indicated on the left and sections excised from the gel separating lipid droplet associated-protein are indicated on the right. 170 UJ TAG FA : Total FA 1... 1.0- d 0.81 0.6- 0.4- d 0.2‘ +N -N -N -N 0 O ' I ' 1 ‘ l 350 400 450 500 550 600 650 700 Wavelength (nm) L0 24h 48h 72b 11’; 250- 150- 100- 80- 60- 50 40 30- 25- 20- 15- 10. Cell \- ~b «‘9 g «0" ". 171 Table 5.2. Fatty acid content of whole organelles (eyespot and lipid droplet) compared to fatty acids bound in triacylglycerols isolated from these organelles. Whole Whole a organelle TAG organelle TAG Fatty acid Eyespot Eyespot Lipid droplet Lipid droplet 16:0 35.5 1 0.67 67.0 1 7.55 32.2 1 1.24 41.8 1 4.33 16:] 3.0 1 0.01 4.5 1 1.08 3.4 1 0.37 3.1 1 0.58 16:2 3.4 1 0.02 3.0 1 0.30 1.2 1 0.15 0.9 1 0.06 16:3 15.7 1 0.18 7.8 1 0.45 6.4 1 3.03 5.1 1 1.70 18:0 3.0 1 0.01 1.9 1 0.50 2.1 1 0.87 1.5 1 0.77 1821,1624 18.4 1 0.39 7.9 1 1.99 27.2 1 1.74 27.3 1 2.86 18:2 10.9 1 0.71 4.7 1 1.77 10.7 1 0.67 10.0 1 1.67 18:31”12 5.6 1 0.09 1.4 1 0.57 7.7 1 1.59 5.9 1 2.62 18:3A9"2"5 3.3 1 0.47 0.8 1 0.41 6.8 1 4.69 2.9 1 0.34 18:4 1.1 1 0.02 1.1 1 0.47 2.3 1 0.92 1.4 1 0.38 a Organelles were isolated from cells grown switched to 0 mM NH: and grown for one day. The data are presented on a % mol basis where two replicates were averaged for eyespot samples and three for lipid droplet samples. The standard deviation is indicated. 172 minimal contamination of thylakoid membranes, the predominant membrane fraction in C. reinhardtii. Analysis of proteins from whole cells, thylakoids, the eyespots, and lipid droplets by SDS-PAGE followed by total protein staining (Figure 5.3C) revealed a pattern for lipid droplets that was distinct from the other protein fractions, with one major protein of approximately 27 kDa, as well as several less abundant proteins, which were Specifically enriched in lipid droplets compared to thylakoids and the eyespots. The lipid droplet proteins were excised from the gel (as indicated in Figure 5.3C) and subjected to mass Spectrometry for protein identification. Proteins identified in the lipid droplet fraction. As summarized in Table 5.3, 259 proteins with two or more unique peptides were identified and fell into a broad range of protein functional groups (Figure 5.4A). Sixteen proteins predicted to be involved in lipid metabolism, including three presumed acyl-CoA synthetases and a predicted lipoxygenase—proteins previously identified as lipid droplet associated in studies of other organisms (Bartz et al. , 2007, Cermelli et al., 2006)—were among those proteins. BTAI, the enzyme responsible for the synthesis of a major extraplastidic membrane lipid diacylglyceryl-N-trimethylhomoserine (Riekhof et al., 2005) was also present. Several predicted components of vesicular trafficking pathways were identified, including subunits of the Coat Protein Complex I and its putative regulator ARF la, as well as other small Rab-type GTPases. Orthologs of these proteins have also been identified in lipid droplet proteomics studies done on different animal species (Bartz et al., 2007, Cermelli et al. , 2006), and recent Drosophila cell RNAi inactivation studies targeting Arj79a (ARFIa ortholog) and the Coat Protein Complex 1 encoding genes revealed a previously 173 Amino acid metabolismls) I (24) Other metabolism Channels (21' c- (3) Oxidative pentose phosphate pathway Cytoskeleton13)l 11(4) Oxidative stress related .1 _ I(2) Photoreceptors Function unknonwn (45)I I(24) Photosynthetic apparatus I(2) Proteases and peptidases Glycolysis (S)I Ion transporting ATPaseS(5)I Kinases and 14-3-3 proteinsl7)I .(7) Protein translocation. chaperones and protein transport Lipid metabolism (16)I . . I(54) Ribosomes. translation Microtubule motor ml and DNA related Mitochondria (26) I I(7) Transporters I(1) Ubiquitin protein modification I(15) Vesicle trafficking Vesicle Trafficking (7) I I (2) Channels I (3) Cytoskeleton Transporters (S)I I(10) Function Unknown Ribosomes, Translation and DNA Related (12) a I(3) Glycolysis I (1) ion transporting ATPases I(3) Klnases and 14-3-3 proteins Protein Translocation, chaperones, and protein transport (2)- proteases and peptidases (1) I I (10) Lipid metabolism Photosynthetic Apparatus (12)- l (8) Mitochondria Photorecptors (1) I oxidative stress related (3) I . (8) Other Metabolism l (2) Oxidative pentose phosphate pathway Figure 5.4. Distribution of lipid droplet associated proteins into functional groups. (A) Functional distribution of all 259 proteins identified with 2 or more unique peptides and (B) proteins (93) with greater than 12 spectral counts. 174 Table 5.3. Complete list of proteins identified in the Chlarnydomonas lipid droplet fraction. 1 . . . . . . Manual annotation and categorization of proteins into functional groups 2Number of Spectral counts or unique peptides identified by LC MS/MS 3Deduced protein molecular weight (kDa) 4Occurrence of identified proteins in C. reinhardtii MS-proteomes of the eyespot (Schmidt et al., 2006), mitochondria (Atteia et al., 2009), or flagellum (Pazour et al., 2005) 5Protein ID and model name for identified proteins (C. reinhardtii genome v. 3.0) v “‘ g"! 59.. 5' E {P '6 Functional category and protein description' g g a g .5 2 g g 3 m." 3': § ‘6 , g i g o g]. 3 Protein ID 6 l u. u. 2 WM 1 Glycine hydroxymethyitransferase 4 3 51845 - 107904 2 alutanb-ganma-semiaidehyde dehydrogenase 12 3 41039 130812 3 glutanine synthetase 12 6 42128 I 133971 4 aspartate semialdehyde dehydrogenase 11 6 39917 148810 5 cobalamin-independent methionine synthase 4 86756 I I 154307 6 Aeetylglutamate kinase-like protein 3 35955 78991 cum 7 Voltage-dependent anion-selective channel protein 16 7 28533 I 127172 8 Porin 21 8 281$ I 146232 9.1mm 9 alpha tubulin 1 47 14 49569 I 128523 10 beta tubulin 2 83 17 49601 I_ 129868 1 1 aclin 23 9 41819 I I 24392 W 12 Truncated Hemoglobin 7 2 15677 159380 13 VFL2 Centrin 2 2 19442 I 159554 14 Calmodulin related protein 6 3 36353 I 183554 15 Calreticulin 2, calcium-binding protein S 3 47311 I 78954 16 predicted protein 18 8 28992 103840 17 sinilar to Hismacro and SEC14 domain proteins 3 2 14867 107313 18 similar to adenylate kinase 8 3 53990 109953 19 similar to aspartate aminotransferase S 3 44429 11 1917 20 predicted protein 11 4 30564 118714 21 predicted protein 11 5 30582 121991 22 similar to cell division inhibitor 18 5 32540 122924 23 glycoside-hydrolase-like protein 4 2 100235 128630 24 Deflagellation inducible protein 4 3 12693 I 131284 25 Flagellar Associated Protein 3 2 67438 I 132213 26 predicted ER protein, similar to disulphide isomerase 4 3 58220 133800 27 s'milar to chloroplast outer envelope protein 7 4 73326 143879 28 Uncharacterized conserved protein 2 2 93303 149656 175 Table 5.3 (cont’d). 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 $8983.88?» 8839'; 61 62 63 65 67 68 69 70 71 72 73 hypothetical protein predicted protein predicted protein predicted protein predicted protein predicted protein predicted protein predicted protein predicted protein predicted protein Flagellar Associated Protein predicted protein Chlamydomonas specific tamiy protein predicted protein mitochondrial trmscription termination factor predicted protein Anlryrin repeat protein TBZIDP1 and HVA22 related protein predicted protein predicted protein Flagellar Associated Protein predicted protein predicted protein predicted protein predicted protein Flagellar Associated Protein predicted protein predicted protein 180660661 Hexokinase Phosphoglycerate Kinase GAP1A glyceraldehyde 3-phosphate dehydrogenase Phosphoglycerate mutase Enolase W Vacuolar ATP synthase subunit 8 plasma membrane-type proton ATPase Vacuolar ATP synthase subunit E vacuolar ATP synthase, srbunit A calcirmr-transporting ATPase, ER Win: Calcium-dependent Protein Kinase 1 cyclic nucleotide dependent protein kinase protein kinase 5‘-AMP-activated protein kinase. gamma subunit 14-3—3 protein 14-3-3 protein Serine/threonine protein kinase W 176 15 13 42 11 nmmawgwwwwnuN-huwwmmoumuhanew $00100) 11 N#O’@Ul “@0001 11 29855 38 1 91 1 521 1 36027 27659 2 1 028 231 30 27063 69974 21621 376% 36056 20380 1 3390 25701 1 3620 20319 19849 24678 42641 22 149 46163 27963 29581 25885 190$ 47143 12745 29524 49015 39742 515$ 55782 1 17266 26226 68378 1 14325 67431 81010 44181 29497 27986 49490 151400 151947 155900 166924 168924 170664 173167 175668 176621 176660 162502 , 163446 183515 164901 185148 165214 165396 166150 166179 166261 169631 192470 192623 193410 194644 194731 195067 79613 123020 132210 140618 161085 83064 126382 182602 190371 54608 77047 128451 131695 153242 184895 185967 187228 93758 Table 5.3 (cont'd). 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 Long—chain acyl-00A synthetases Long-chain acyl-00A synthetase Cycloartenol synthase coclauine N-methyltiansferase Long-chain acyl-00A synthetases (AMP-forming) VTE3 MPBQ/MSBQ methyltransferase Lipoxygenase pyruvate-tormate lyase Esterase/Iipaselthioesterase Predicted phosphate acyltransferase 4'-phosphopantetheinyt transferase Phospholipase DfTransphosphatidylase TEF21 phosphoinositide phosphatase 3-beta hyrkoxysteroid dehydrogenasefisomerase Betaine lipid synthase Predicted phosphate acyltransferase, W Cytoplasmic dynein 1b heavy chain In ! I . Mitochond' rial substrate cam'er MPC1 Mitochondrial Phosphate Carrier 1 UQ:Cth oxidoreductase 50 kDa core 1 subunit NADqubiqu'none oxidoreductase sibunit 10 mitochond' rial F1F0 ATP synthase 36.3 kDa protein predicted mitochondrial substrate carrier mrtochond' rial F1F0 ATP synthase 31.2 kDa protein NADH:ubiquinone oxidoreductase N09 subuni NADqubiqu'none oxidoreductase 16 kDa subunit mitochond’ rial F1F0 ATP synthase 14.3 kDa protein NADqubiquinone oxidoreductase 49 kDa subunit Mitochond' rial F1F0 ATP synthase. delta subunit cytochrome c oxidase submit NADqubiquinone oxrdored' uctase 39 kDa subunit mitochond' rial F1F0 ATP synthase 13.3 kDa protein Mitochondna' ' lcytoctrorne c oxidase 13 kD sibun'l Cytochrome c oxidase subunit ll. protein llb mitochond' rial substrate carrier protein mitochondrial substrate carrier protein mitochondrial F1F0 ATP synthase 19.5 kDa protein NADqubiquinone citidoreductase 18 kDa subunit F1F0 ATP synthase gamma subunit ATP synthase, alpha subunit UQ:Cth oxidoreductase 14 kDa sibunit beta subunit ot mitochondrial ATP synthase mitochondrial F1F0 ATP synthase, 60.6 kDa protein museum NAB-dependent epimeraseldehydratase uridine 5‘- monophosphate synthase glycosyl transferase, group 1 S-Adenosyl homocysteine hydrolase 177 1 30 62 74 78 92 13 11 13 231 14 11 12 12 18 160 5.: N cwwbwawwouw b 0001 10 21 18 8 : Add—s #- 000163000116:- 0907.5th NUNUIUI (D 4.6 501 UIUNNUIUWwa .6 #0) 17 19 14 10875 55555 85316 41223 72232 37446 1 18099 91 132 271 14 40174 91780 245% 50595 41515 75764 23923 481450 28548 37422 55037 18104 39710 34000 32124 15253 14307 52591 21154 11665 43637 16166 19010 17232 31616 25861 22185 17995 35077 61497 14024 61804 63145 36459 50921 46161 52701 111160 113915 116558 119132 123147 129760 144677 146801 160378 189038 189764 190403 190968 58501 77062 98450 24009 118965 135199 138803 139850 152682 159938 182740 182980 184222 184815 185013 185200 185375 186185 186501 187882 190125 191516 191596 192157 193762 194183 76602 77031 78348 78831 111778 116267 122195 129593 Table 5.3 (cont'd). 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165 166 cytochrome b5 protein gamma carbonic anhydrase type-ll calcium-dependent NADH dehydrogenase Superoxide dismutase. MAD-dependent malate dehydrogenase Apoferredoxin NADH-cytochrome b5 reductase sinilar to glycosyl hydrolase succinate dehydrogenase soluble inorganic pyrophosphatase S-adenosylmethionine synthetase chitinase—related protein UDP-glucose dehydrogenase Malate dehydrogenase. isocitrate lyase gamma carbonic anhydrase putative quinone oxireductase Short chain dehydogenase/reductase family protein nucleoside diphosphate kinase-lire Malate dehydrogenase Transaldolase 6-phosphogluconate dehydrogenase 2W 2-cys peroxiredoxin, chloroplastic glutathione S-transterase glutathione-S-transferase Sinilar to peroxiredoxin 4 (Bos taurus) EM! COP2 chlamyopsin retinal binding protein phototropin W Ferredoxin-NADP (+) reductase. CFO ATP synthase subunit ll precursor Photosystem l protein PsaD Chlorophyll A-B binding protein PSBO Oxygen-evolving enhancer protein 1 ll Chlorophyll A-B binding protein Chlorophyll A-B binding protein LHCA6 light-harvesting protein of photosystem I chloroplast ATP synthase delta chain LCHAS light-harvesting protein of photosystem I LHCA8 Photosystem II reaction center protein PsbP PSBQ Chlorophyll A-B binding protein Chlorophyll A-B binding protein Rieske ferredoxin PSAH Subunit H of photosystem I 178 U'lmNhflmm 137 WM 12 21 30 32 14 10 16 17 19 73 10 10 37 59 10 14 15 29 10 46 16 0017019301“ (0 1 mObNN 1 (DON NWQONN§ 0004501 14 ("#030203 a O N§bwb00101NUlw 14988 31 1 50 66955 25898 32267 13442 31 135 74848 49697 22157 42566 22350 52707 36584 45732 24274 21427 24604 16523 77644 42419 53348 25944 16037 23941 21624 81450 38231 1 6554 20891 22827 30697 28670 27764 23977 2821 1 25904 25882 21807 23883 27549 14991 14134 131692 132797 133334 135320 158129 159161 163751 169029 172283 174103 182408 184328 185081 190455 191668 195016 24039 55666 58944 60444 133861 146574 15891 1 142363 154723 193661 53816 164843 183965 105589 105641 120177 128836 130316 130414 131 156 131602 132678 133575 134203 148057 153656 174723 178631 182093 182959 Table 5.3 (cont’d) 167 168 169 170 171 172 173 174 175 176 177 178 179 180 181 182 183 184 185 186 187 188 189 190 191 192 193 194 195 196 197 198 199 201 202 203 205 207 208 209 210 211 212 213 Chlorophyll A-B binding protein LHCBMG chloropyll a-b binding protein of LHCll LHCB4 chlorophyll a-b binding protein Chlorophyll A-B binding protein Chlorophyll A-B binding piotein Chlorophyll A-B binding protein PSBR 10 kDa photosystem II polypeptide W Predicted serine protease Peptidase M14. carboxypeptidase A LA HSP70-like protein Heat shock protein 90A similar to Tic32-3 Translocon-associated complex alpha subunit Heat shock protein 70A SECS1-alpha subunit of ER-translocon Peptidyl-prolyl cis-trans isomerase, cyclophilin type W Histone H4 Histone H2A Ribosomal protein L10E Histone-lire bacterial DNA-binding protein R'bosomal protein 86 Ribosomal protein L7Ae Receptor of activated protein kinase C 1 RNA-binding protein Ribosomal protein S17e ATP-dependent RNA helicase Ribosomal protein $9 608 ribosomal protein L-11 Ribosomal protein 608 Ribosomal protein S4 Rbowmal protein L356 Ribosomal protein L13 eukaryotic translation elongation factor 1 alpha 2 Ribosomal protein 83 Ribosomal protein 810 Ribosomal protein L9 eukaryotic initiation factor Ribosomal protein S19 Ribosomal protein 824 Ribosomal protein L186 Ribosomal protein L14 elongation factor EF-3 Ribosomal protein L18a Ribosomal protein 810 Ribosomal protein 327 isofonn A Ribosomal protein L28 Ribosomal protein L276 179 19 10 10 16 19 mfifllflflm NWNUN‘U (JINUlbUthUOUIQUéNUl @b‘lh # .5 N wwwwwwmonmwwau 26670 26944 29921 27363 28685 26205 13025 85142 281% 72477 80667 38224 27049 71 198 52488 22660 11440 13668 26534 8949 28418 27091 35127 25272 15880 63436 15916 19469 9948 21154 12040 24466 50807 25830 13290 21501 24231 16980 15105 23958 15324 115293 21354 19291 9497 19480 16373 184070 184490 18481 0 186064 24036\ 78552 98021 147894 184660 133859 1381 17 145585 185608 185673 188207 118846 100020 100147 103135 104082 104348 105289 105734 107394 108082 111269 113739 116329 118268 122980 127247 129809 132905 134051 139083 139831 140593 141798 143072 144684 145271 145770 146844 149993 150644 153674 155455 Table 5.3 (cont’d) 214 215 216 217 218 219 220 221 222 223 224 225 226 227 228 229 230 231 232 233 234 235 236 237 238 239 240 241 242 243 245 247 248 249 251 252 254 255 256 257 258 259 Ribosomal protein S 18 Ribosomal protein L23 Histone H26 variait Ribosomal protein L136 Acidic ribosomal protein P0 Ribosomal protein 83a, Ribosomal protein L6 glycine rich RNA binding protein Ribosomal protein S4 eukaryotic initiation factor 4A-Iike protein Ribosomal protein L30 Ribosomal protein 65 Ribosomal protein L22 eukaryotic initiation factor Ribosomal protein 814 Ribosomal protein L21 EFGZ Elongation Factor 2 Ribosomal protein L27 Ribosomal protein L24E Ribosomal protein 65 Ribosomal protein S156 Ribosomal protein 88E Glycyl-tRNA synthetase and related 1m similar to 1m phosphate transporter Adenine nucleotide translocator Sunni warmer swerfamily putative 2-oxoglutaratelmalate trmsporter Nitrate transporter Major facfl'tator superfamily protein plastidic ADP/ATP translocase Ubiquitin-conjugating enzyme E2 W Clathrin propeller, N-terminal. Ras GTPase RA823 smal Rab-related GTPase alpha-COP small Rab-related GTPase COPBZ Beta'-COP small Art-related GTPase ARFA1a similar to VAMP-associated protein Dynamin GTPase effector COPE1 Epsilon-COP small Rab-related GTPase COPG1 Gamma-COP small Rab-related GTPase Sar-type small GTPase Dynamin. Dynamin central region 180 11 20 14 16 10 10 13 26 10 26 S3 19 32 111 51 22 15 QANNbO’hQA-‘Qtflh‘mm a 5 10090901100) d bNdVNmQhOb a #0) 12 11 17562 14916 16537 20710 34557 29314 24355 15766 29467 47037 27559 21690 14444 23079 16284 18300 94193 15248 13636 30273 14821 25176 76729 31 660 3351 1 33431 481 93 57394 62295 16038 190068 21959 19725 1381 16 23594 107719 20569 27963 60945 31578 22581 96947 22103 21847 67594 155649 159282 160509 162845 164097 168484 183518 184151 188837 188942 190273 191776 1951 31 24172 24344 24354 24524 260 44781 56455 59755 76860 82920 116544 138185 169636 178501 192090 193659 195230 187409 1 12083 1 16488 126840 143349 148836 154280 183074 183897 186497 190416 60490 78205 81259 82091 98432 unknown role for these proteins in lipid droplet formation and utilization (Beller er al., 2008, Guo et al., 2008). The presence of presumed orthologs in C. reinhardtii lipid droplets suggests the possibility of a similar role in algal lipid droplet biogenesis. Other functional groups which were represented by many proteins included 54 predicted ribosomal and translation related proteins, 26 predicted mitochondrial proteins, and 24 predicted proteins of the photosynthetic apparatus. Ribosomal and mitochondrial proteins have been identified in lipid droplet proteomic studies of different animal species (Cermelli er al., 2006, Walther and Farese, Jr., 2009) and plant oil bodies (in plants from mitochondria only) (Katavic er al., 2006), and could be common contaminating proteins in lipid droplet preparations, or alternatively could indicate the presence of ribosomal- /mitochondrial-lipid droplet physical interactions, as has been previously postulated (Cermelli er a1. , 2006). The identification of 24 photosynthetic apparatus proteins was surprising, given the near absence of chlorophyll in purified lipid droplets, and it is likely that these hydrophobic apo-proteins are contaminating artifacts that become associated during the isolation procedure; however, the most abundant stromal enzyme, ribulose- 1,5-bisphosphate carboxylase/oxygenase, was not detected. A more stringent analysis of 93 proteins (identified by more than 12 spectral counts) resulted in 6 ~38 % decrease in the number of lipid metabolism proteins retained in the “identified” group (from 16 to 10), whereas relative decreases in the more predominant contaminating groups were generally higher (~78 % for ribosome/DNA/translation related, ~69 % for mitochondrial and ~50 % for photosynthetic apparatus proteins—see Figure 5.4B). In addition, 45 proteins of unknown function were identified. Based on spectral counts for all unique peptides for a given protein (Table 5.3), one of these was by a factor of lo the most 181 abundant protein in gel slice 9 (Figure 5.3C) which also contained the most abundant protein in the lipid droplet fraction based on staining. Its calculated molecular mass is 28 kDa which approximately corresponded to its position in the gel (Figure 5.3C). Therefore, this protein was presumed to be the most abundant protein in the lipid dr0plet fraction and was tentatively designated major lipid droplet protein (MLDP—protein ID: 192823 in the C. reinhardtii v. 3.0 database http://genome.jgi- p psf.org/Chlre3/Chlre3.home.html;_NCBl accession number XP_001697668). Notably, the second most abundant protein based on spectral counts was BTAl, the enzyme catalyzing the biosynthesis of betaine lipid. Both proteins were also identified in study of C. reinhardtii lipid droplets conducted in parallel, but dismissed as contamination (Wang et al., 2009). MLDP suppression affects lipid dr0plet size. Steady-state mRNA abundance of MLDP was determined by northern blotting and quantitative real-time RT—PCR in 10 mM N- grown cells, and cells deprived of N for 24, 48 and/or 72 h as shown in Figure 5. The widely used C. reinhardtii control gene RACK I (CBLP) was used as an internal control for RT-PCR. Analysis of apparent fold change in RACK 1 expression by the 2"“ method (see Materials and Methods) indicated that changes in its expression based on total RNA input to the reverse transcriptase reaction were minor if any (~l .5 fold increases at 48 and 72 h -N). Previous studies have shown that during N-deprivation in C. reinhardtii, the amount of chloroplast and cytoplasmic ribosomes and total RNA content decrease (Martin 6101., 1976). While this can potentially add an additional level of difficulty in identifying a suitable control gene in these experiments, the degree to which this 182 contributes to the apparent minor fold-changes in expression for RACK 1 is unknown. With the above in mind, RACK 1 was deemed a suitable control gene for these experiments, with the caveat that it may slightly underestimate the fold-changes for genes induced by 48 or 72 h N—deprivation, and vice versa for down-regulated transcripts. When normalized to RACK 1 , MLDP mRN A showed a 16-fold increase in abundance after 24 h of N—deprivation and continued to be abundant even after 72 h of N- deprivation. Moreover, MLDP mRN A abundance qualitatively corresponded during a time course of N-deprivation with the accumulation of TAG (Figure 5.1A). To begin the functional analysis of MLDP in C. reinhardtii MLDP RN Ai lines were generated. For this purpose, a plasmid carrying an expression cassette which is inducible by N- deprivation, pNITPROl , was constructed based on the nitrate reductase (NI T I ) promoter. The pNITPROl vector was used to drive the expression of an MLDP genomic sense- cDNA antisense RNAi hairpin (Figure 5.6A). Zeocin-resistant transformants were screened for MLDP mRNA reduction at 48 or 72 h N-deprivation. Of 64 lines screened, 9 lines were identified with a range of MLDP transcript reduction between 20 and 60% of empty vector controls. The four strongest MLDP-RNAi lines (A l , C7, E9, and G1 1) showed a consistent 55-60% reduction of MLDP transcript measured after 72 h N- deprivation from three independently grown cultures (Figure 5.6B). The MLDP-RNAi lines were analyzed for possible lipid droplet morphological phenotypes by fluorescence microscopy analysis of Nile-Red-stained cells deprived of N for 3 days. An apparent increase in the average size of lipid droplets in the MLDP-RNAi lines was observed (Figure 5.7). The apparent individual diameters of a population of greater than 200 random and in focus lipid droplets per RNAi or control lines were 183 B 20 1 «$529 6““‘0’6 g ' «9 x‘ \9 x‘ '2 16: 3 .. 12 a: MLDP 5 - e. Z 8 J 1: . E 4 - i g ., rRNA 0 . + N 24h 48h 72h -N Figure 5.5. MLDP mRNA is induced following nitrogen deprivation. (A) Northern blot analysis of MDP, where 5 pg total RNA isolated from l0 mM NH4Cl grown cells (10 mM N) and cells deprived of N for 2 days (-N 2 d) was processed for northern blotting. Total RNA was stained with methylene blue and shown as a loading control (rRNA). Two biological replicates of this experiment are shown (panels 1 and 2). (B) Relative abundance of MLDP mRNA normalized to RA CK I (CBLP) in wild-type dw15 cells grown in medium with 10 mM NHf (+N), or dw15 cells switched to and grown in medium with no NHf {-N) for 24, 48, or 72 h measured by QRT-PCR. Data are expressed as a fold change compared to the +N grth condition which are given a value of l. Standard deviation of three biological repeats, each done in triplicate is shown. 184 RBCSZ 3’UTR NIT1Pr0 ll— ...5’UTR 1 91 ‘_ ‘ .nJJIlllllllll] i .' ~ ’ 91 310 422 "301R .3 N Rel mRNA Abundance w .o .o ‘5 °.° A O EV1 EV2 A1 C7 E9 G11 Figure 5.6 Generation of MLDP RNAi-mediated knockdown strains. (A) Diagram showing the MLDP-RNAi construct designed with the NIT 1 inducible promoter (Pro) of vector pNITPROl driving the expression of an MLDP genomic inverted repeat (numbers indicate basepairs from predicted S’UTR), and the predicted transcribed RNA hairpin product. (B) QRT—PCR analysis of MLDP mRNA levels in two empty vector control lines (EVl and EV2), and four MLDP-RNAi lines isolated from four independent transformation events (A1, C7, E9, 61 1). Data are represented as percent change compared to RV] and normalized to RACK] as in (A). Three replicates were averaged and standard deviation is shown. 185 > L0 Diameter (pm) OJ E2511) E2613 (2213) (£56) (2E299) (2114} A1... Figure 5.7. MLDP RNAi knockdown lines exhibit a lipid droplet morphological phenotype. (A) Analysis of apparent lipid droplet diameter in empty vector controls and MLDP-RNAi lines shown in Figure 5.5. The average and standard deviation are shown for n (indicated in parentheses below each bar) lipid droplets counted, and cell lines are marked with an asterisk, for which P < 0.01 when compared to EV] in a two-tailed Student’s t-test. (B) Fluorescent micrographs of representative cells of lines EVI and Al stained for lipid droplets with Nile Red. 03 EV1 186 measured after three days of N-deprivation (Figure 5.7A). An approximate 40% increase in the average lipid droplet diameter was determined in the MLDP-RNAi lines. MLDP has orthologs only in green algae. The newly discovered MLPD protein has few orthologs, none of which has been functionally characterized. When searched against the National Center for Biological Information database of non-redundant proteins using BLAST (Altschul er al., 1997) no sequence with significant similarity is identified. However, several genome projects of green algal species are currently underway and searching against green algal genomes at the respective websites, potential orthologs with high sequence similarity were identified in Chlorella sp. NC64A, Chlorella vulgaris and Volvox carterr'. No discemable ortholog was present in available diatom or red algal genomes or the genomes of Osrreococcus species, which belong to a green algal lineage distinct from C. reinhardtii. Apparently, MLDP is a rare protein in nature as we currently know it at the molecular level and presumably arose during the evolution of green algae. As such it also has no discernible domains to which functions have been assigned. Grand average of hydropathy (GRAVY) indices and Kyte-Doolittle (Kyte and Doolittle, 1982) hydropathy plots were generated for MLDP, as well as for mouse ADRP and Perilipin, and Arabidopsis Oleosin l (OLEl) (Figure 5.8 A-D). All four proteins were distinct with respect to extent and region of hydrophobicity, with the most hydrophobic region for MLDP occurring between amino acid residues ~l 60-215. The proteins ranked according to their GRAVY index (highest to lowest) are: MLDP (0.l l), OLE] (0.17), 187 Figure 5.8. Hydropathicity plots of MLDP and other known lipid droplet binding proteins. (A-D) Hydropathicity plots for the proteins (followed by NCBI accession numbers in brackets) MLDP [XP_001697668] (A), mouse ADRP [NP_031434] (B), mouse Perilipin [NP_783571] (C), and Arabidopsis OLE] [NP_l94244] (D). The value G in each graph is the grand average of hydropathy value (GRAVY) for each protein and was calculated using the GRAVY calculator program at http://gravyjaborfi’ustdel. 188 35- 236:041 15. 05. 415. 445 415. 3.5 v v r ' 0 50 100 150 200 250 C3 ' 0 100 200 300 400 35- 2.5- = -0.40 [3 ' 0 100 200 300 400 500 0 50 100 150 189 ADRP (028), and Perilipin (-.40). Thus, MLDP is a very hydrophobic protein although it lacks a specific hydrophobic core characteristic for oleosin. Discussion The potential of microalgae to provide sustainable feedstock for biofuel production has led to renewed interest in the biology of this diverse group of organisms (Hu et al., 2008). While C. reinhardtii is unlikely to become the microalgae of choice for the production of biofuels, it is a well established genetic and genomic model for the study of unicellular green algae (Merchant er al., 2007). However, very little is known about lipid metabolism or the cell biology of lipid droplets and its regulation in this organism. The time course and conditions for the induction of oil accumulation described here (Figs. 5.], 5.2) and the detailed analysis of lipid droplets provides the basis for a molecular analysis of the mechanisms of TAG accumulation in C. reinhardtii and possibly unicellular green algae in general. As a logical first step, we focused on the TAG-storing compartment itself, the lipid droplet. Its fatty acid and pigment composition is clearly distinct from that of other organelles (Figure 5.3). This distinct composition and the very low chlorophyll content in view of the predominance of thylakoid membranes in the cell suggested that the obtained oil droplet preparation was only slightly contaminated with thylakoid membranes, the largest membrane system in the cell. However, we cannot rule out some level of contamination with ER—derived microsomes especially since lipid droplets are generally thought to be derived from the ER. An increase in the total amount of fatty acids in N- deprived cells indicated that the biosynthesis of TAGS is not merely due to conversion of membrane lipids but the result of de novo synthesis. The specific fatty acid composition 190 of lipid droplet lipids tended towards shorter and more unsaturated fatty acids suggesting that newly synthesized fatty acids were quickly incorporated into TAGs before fatty acid modification by desaturases or elongases can occur; alternatively, reduced activities of these fatty acid metabolic enzymes could account for this observation. Lipid droplets in different organisms are increasingly recognized as dynamic organelles that provide important nodes of cellular metabolic networks and participate in intracellular lipid trafficking (e.g. Guo et al., 2008). In algae, induction of lipid droplet formation is intricately regulated by nutrient deprivation and other abiotic stresses that lead to a cessation of cell division or an over-reduction of the photosynthetic electron transport chain (Hu et al., 2008). In this study, we sought to identify the proteins associated with lipid droplets to determine factors involved in the biogenesis or turnover of this organelle in C. reinhardtii. Aside from those of unknown function, of particular interest are lipid metabolic enzymes and proteins involved in vesicular transport Intriguingly, the betaine lipid (diacylglyceryl-MMN-trimethylhomoserine) biosynthetic enzyme, BTAI (Riekhof er al., 2005), had the second highest spectral count of proteins in the lipid droplet fraction. This protein was also identified as one of few in the lipid droplet fraction in a parallel study, but considered a contamination (Wang et al., 2009). This parallel study concluded that no protein was specifically associated with the respective lipid droplet fi’action prepared and treated differently than in this study. Here, a number of predicted acyl-CoA synthetases and acyltransferases were present as well that could be involved in the biosynthesis of TAGs or the transfer of acyl groups between membrane lipid and neutral lipid pools. In animal cells, enzymes involved in phosphatidylcholine biosynthesis have been found critical for lipid droplet 191 formation and turnover. Lipid dr0plets in animal cells are surrounded by a monolayer of phospholipids and the availability of those phospholipids affects lipid droplet size. When the rate-limiting enzyme of phosphatidylcholine biosynthesis was inactivated in Drosophila cells, lipid droplets increased in size presumably because the larger droplets have a lower surface area to volume ratio and require less phospholipid to form the phospholipid monolayer (Guo et al. , 2008). As C. reinhardtii lacks phosphatidylcholine, it seems plausible that its role in the formation of lipid droplets is taken by betaine lipid, which has similar properties and is thought to functionally substitute for phosphatidylcholine in this green alga (Sato and Murata, 199]). A rich complement of proteins predicted to be involved in vesicular trafficking including COP] homologs, Rab GTPase, or ARF related GTPase were also identified in the proteomics data set. Repression of the respective orthologs in Drosophila cells was shown to affect lipid droplet formation and dynamics and TAG utilization (Guo et al., 2008). Our understanding of the interaction of lipid droplets with the vesicular transport machinery is only rudimentary at this time. The presence of the respective proteins in C. reinhardtii follows a common theme also observed for animal cells and provides an opportunity to study the role of the vesicular transport machinery in lipid droplet formation in a unicellular model organism. However, the specific association of these proteins with lipid droplets in C. reinhardtii remains to be confirmed. The identification of proteins in the ribosome-ltranslation-lDNA-related, mitochondria, and photosynthetic apparatus functional groups suggest a basic level of contamination in our lipid droplet protein fraction. While more stringent filtering of the proteomics data set to include proteins with more than 12 spectral counts resulted in a 192 larger decrease in the relative proportion of these apparent contaminating functional groups, when compared to lipid metabolism proteins (Figure 5.4, Table 5.3), the significance of such analyses should be regarded as circumstantial given the potential for protein size bias in the method. Comparing the analysis of the lipid droplet proteins presented here with proteomic studies of the eyespot (Schmidt etal., 2006), flagella (Pazour er al., 2005), and mitochondria (Atteia et al., 2009) (Table 5.3) reveals that some degree of contamination with proteins from specific functional groups is common (e.g. ribosome/nanslation-IDNA-related, mitochondria and photosynthetic apparatus proteins found in the eyespot study, or ribosome contamination in the mitochondrial and flagellar studies). As such, apparent contamination of subcellular preparations in C. reinhardtii seems to be a common challenge. To begin the functional analysis of lipid droplet associated proteins in C. reinhardtii we focused on the most abundant protein in the lipid droplet fraction based on Coomassie Brilliant Blue staining (Figure 5.3C) and spectral counts (Table 5.3). The identity of this protein was deduced by mass spectrometric analysis of all proteins in the respective gel slice and MLDP was identified based on lO-fold overabundance of spectral counts for its peptides. Repeated attempts to localize an MLDP-Green Fluorescent Protein (GFP) fusion protein to the lipid droplets failed because none of the tested constructs expressed in C. reinhardtii. Unlike plants, C. reinhardtii does not seem to have an ortholog of oleosins, distinct structural proteins surrounding the lipid bodies in plant oil seeds (Huang, 1992, Siloto er al., 2006). Interestingly, oleosins are also absent from oil-rich mesocarp tissues of plants like oil palm or olives and their presence may be related to special needs of seeds. Indeed, reduced oleosin content leads to delayed 193 germination and reduced freezing tolerance (Shimada er al., 2008, Siloto et al., 2006). At the cellular level, a reduction in oleosin caused an increase in oil body size. Likewise, when MLDP was inactivated in C. reinhardtii, an increase in lipid droplet size was observed (Figure 5.7). Despite an extensive screening of transgenic lines expression of the MLDP gene could only be reduced approximately 60% in the best RNAi lines leading to a moderate but statistically significant increase in oil droplet size in these lines. Because MLDP lacks any features that readily classifies it as a protein with known 1"" catalytic activity, it may function as a structural protein affecting lipid droplet dynamics and size in C. reinhardtii. An increase in lipid droplet size has been shown in different systems to delay turnover of TAGs. We measured TAG levels (at 3 d —N) and TAG ls turnover in these lines following the resupply of N in the medium but could not detect a statistically significant difference in the levels of TAG or rate of TAG metabolization in these lines compared to the empty vector control lines. It seemed possible that the observed moderate change in lipid droplet size would lead to only a small change in the rate of TAG metabolization, too small for it to be detected within the statistical limitations of our analytical method. This does not rule out the possibility that MLDP is important in preventing lipid droplet fusion to provide a larger surface area per volume ratio to increase the accessibility of enzymes metabolizing TAGS. Intriguingly, lipid droplets isolated from N-deprived cells of the green alga Eremosphaera viridis (Chlorophyceae) were found to have 26, and 28 kDa associated proteins (V echtel er al., 1992); while these masses are similar to that of MLDP, the study was conducted prior to the development of facile protein identification by mass spectrometric methods, and as such, the identity of the E. viridis lipid droplet proteins remains unknown. 194 The newly discovered MLDP is a rare protein in nature unique to green algae. The expression of its gene is strictly limited to conditions favoring TAG biosynthesis, providing indirect evidence for a role of MLDP in lipid droplet formation or maintenance. These properties potentially make MLDP a marker for lipid droplets and TAG accumulation. References Abramoff, M. D., Magelhaes, P. J ., and Ram, S. J. (2004) Image processing with Image]. Biophotonics intern. l 1, 36—42. Allen, M. D., Kropat, J ., Tottey, S., Del Campo, J. A., and Merchant, S. S. 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Van Donk, 13., Lurling, M., Hessen, D. 0., and Lokhorst, G. M. (1997) Altered cell wall morphology in nutrient-deficient phytoplankton and its impact on grazers. Limnology and Oceanography 42, 357-364. Vechtel, B., Eichenberger, W., and Ruppel, H. G. (1992) Lipid Bodies in Eremosphaera viridis De Bary (Chlorophyceae). Plant Cell Physiol 33, 41-48. Walther, T. C. and Farese, R. V., Jr. (2009) The life of lipid droplets. Biochim. Biophys. Acta 1791, 459-466. Wang, Z. T., Ullrich, N., 100, S., Waffenschmidt, S., and Goodenough, U. (2009) Algal lipid bodies: stress induction, purification, and biochemical characterization in wild-type and starchless Chlarnydomonas reinhardtii. Eukaryot. Cell 8, 1856-1868. 198 Chapter 6 Conclusions and perspectives 199 Conclusions and perspectives During the past five years in which the research for this dissertation was conducted, tremendous advances have been made in our understanding of glycerolipid metabolism in plants. Lipid metabolic pathways are tightly regulated during normal growth and development, but can be highly modulated in response to environmental stress. A number of new enzymes and components of lipid trafficking pathways have been identified, resulting in the emergence of a clearer overall picture of the interplay between chloroplast and endoplasmic reticulum (ER) localized glycerolipid assembly pathways (Benning, 2009, Wallis and Browse, 2010). The regulation of chloroplast galactolipid metabolism has been a major focus in the field, not only due to the predominance of these lipids in plant leaf membranes, but also because of their essential role in providing a matrix for the thylakoid-localized photosynthetic machinery (Diirmann and Benning, 2002). There are two well-studied pathways for galactolipid biosynthesis in Arabidopsis: a constitutive and an alternative induced pathway activated by stresses such as phosphate deprivation. A major question that existed at the onset of this work (and one that still stands) is: what are the factors required for activating the alternative galactolipid pathway during phosphate deprivation? DGSI is not directly involved in the regulation of galactolipid biosynthesis in plants. The work described in Chapter 2 was aimed at answering the above question. Suppressor mutants in the dgdl mutant background, which is deficient in the constitutive pathway for DGDG biosynthesis, were identified by Changcheng Xu and colleagues in the Benning lab (Xu et al., 2003). The suppressors fell into two classes. First the tgd 200 mutants, which constitutively accumulate oligogalactolipids through the activation of a galactolipid:galactolipid galactosyltransferase (GGGT-further discussed below). Further detailed analyses of the tgdl/2/3/4 mutants revealed an essential role for the TGDl/2/3/4 proteins in the chloroplast to ER lipid trafficking pathway (Awai et al., 2006, Benning, 2009, Lu et al., 2007, Xu et al., 2003, Xu et al., 2008). The second suppressor class consisted of the dgdl suppressor I (dgsl-1) mutant (Xu et al., 2003). The gene affected by the dgsl-l mutation was found to encode 6 ~70 kDa protein of unknown function, DGSI , in which the mutation caused a D457N mutation. Work described in Chapter 2 describes the detailed characterization of DGSI through the analysis of the dgsl-l mutant as well as a null insertion allele dgsl-2. The DGSI protein was found to be localized to the mitochondrial outer membrane in a high MR complex. Importantly, the dgsl-I mutation was found to behave semi-donrinantly, and the phenotypes observed in this mutant background including activation of DGDl -independent galactolipid biosynthesis, increased hydrogen peroxide levels, and lowered alternative oxidase protein levels were not observed in the dgsl-2 null allele. Moreover, the alternative galactolipid pathway in dgd1 dgsl-2 double mutants was not affected when compared to dgd1 plants, and no clear phenotypes were established for dgsl-2 mutants. The dgd1 dgsl-l double mutant also showed a larger increase in DGDG accumulation during phosphate deprivation when compared to the dgdl control, indicating that dgsl-1 suppresses dgdl by activating a galactolipid pathway that is distinct from the phosphate regulated one. With these observations, it became evident that DGSI is not directly involved in the regulation of galactolipid biosynthesis in plants, and the activation observed in the dgsl -1 background is a secondary effect, which is commonly observed in dominant alleles (Veitia, 2007). 201 If DGSl is not involved in regulating galactolipid metabolism, then what is its function? A lack of observable phenotypes in the dgsl-2 null allele compounds the difficulty in determining the function of this unknown protein. DGSl has orthologs in other plants, algae, fungi, protists and yeast. The Saccharamyces cerevisae ortholog, NCA2, has been studied as a mutant, but only shows phenotypes as a double mutant with the gene encoding NCA3 (Carnougrand et al. , 1995). Lack of phenotypes in both Arabidopsis and yeast mutants in shared DGSI orthologs could be due to conditional importance of these proteins, where the right stress condition has not been investigated. Though DGSland its orthologs have no apparent paralogs within any species, non- homologous redundant proteins that have evolved through convergent evolution to have the same function as DGSl-like proteins could also result in a lack of observable phenotypes. While not found in all organisms, DGSI orthologs have been maintained in genomes of highly diverse species, suggesting a potential for conditional importance of DGSl. Future work on DGSI should focus on the null dgsl-2 allele, possibly running it through a gauntlet of stress conditions to see if any phenotype can be associated with the loss of DGSI. Furthermore, with ever-expanding genomic resources available, the use of comparative genomics approaches (Hanson et al., 2010) may assist in developing hypothesis-driven research in identifying the function of DGSI. GGGT is encoded by the SFR2 gene in Arabidopsis and galactolipid remodeling in the chloroplast envelope is essential for freezing tolerance. Analysis of galactolipids in dgd1 dgsl-I plants during phosphate deprivation revealed the condition-specific accumulation of oligogalactolipids (Chapter 2). This was 202 similar to the above-described observations in the tgd class of dgdl suppressor mutants. In both cases, the oligogalactolipids formed in these mutants were from the presumed activity of GGGT, an enzyme capable of transgalactosylation between galactolipids, which through processive activity is able to synthesize the oligogalactolipids tri- and tetra-galactosyldiacylglyerol. GGGT remained unidentified in molecular/ genetic terms at the onset of this study (Benning and Ohta, 2005). Chapter 3 describes a reverse genetics approach that was successful in identifying the GGGT-encoding gene. Importantly, this work revealed GGGT to be encoded by a previously described gene in Arabidopsis, SENSITIVE T 0 FREEZING 2 (SFR2). SFR2 was first identified through a forward genetic screen looking for mutants defective in acquired freezing tolerance, and found to encode a glycosyl hydrolase family 1 (GH-l) enzyme of unknown function (Thorlby et al., 2004). Work in Chapter 3 shows that sfi'2 null mutants are unable to produce oligogalactolipids, which are formed at the expense of monogalactolipid (MGDG) during freezing stress in wild-type plants. In addition, triacylglycerol accumulates during freezing through the acylation of diacyl glycerol produced by the GGGT reaction. The SF R2 protein was found to be highly active when expressed in yeast, and was sufficient to catalyze GGGT activity (discussed in detail in Chapters 1 and 3). Considering this data, along with the wealth of information generated in understanding the mechanisms of freezing damage in plants (Steponkus, 1984, Thomashow, 1999), a model for the role of SFR2 in membrane protection during freezing was formed. During freezing, membranes are the site of damage due to cellular dehydration. SFR2 (GGGT) remodels the chloroplast outer envelope lipid composition by converting non-bilayer forming MGDG to bilayer forming digalactolipid and oligogalactolipids in order to prevent membrane 203 fusion in a dehydrated state. Triacylglycerol formed during freezing stress may act as a means of removing membrane lipids and reducing the effective bilayer surface area of a given membrane in order to cope with a shrinking cell volume during dehydration. While the identification of GGGT and its role in freezing tolerance are a significant advance in understanding galactolipid biosynthesis and its role in plant stress responses, many questions about GGGT/ SF R2 remain. First among these is whether or not the role of GGGT/SF R2 is restricted to freezing tolerance. Orthologs of SFR2 are present in plants with no freezing tolerance (e.g. maize), suggesting that SFR2 may play a more general role in lipid remodeling during severe water deficit—drought and freezing tolerance responses in plants have long been known to have significant overlap (Thomashow, 1999). Analysis of the 31?? mutants described in Chapter 3 for sensitivity to drought or high salinity will be a straightforward approach in addressing this question. In addition, the analysis of sfr2 tng/2/3/4 double mutants could reveal the significance of the constitutive activation of GGGT in the tgd class of mutants. Importantly, the mode of activation of GGGT/SFR2 activity is still unknown. The SFR2 protein is constitutively present and not regulated by cold acclimation in Arabidopsis leaves (Thorlby et al., 2004), and GGGT can be activated by a number of hyperosmotic treatments of plant leaves (Chapter 3). All of this indicates a post- translational mode of activating the enzyme during hyperosmotic stress. One potential mode of activation could occur through post-translational modification (PTM), e. g. reversible protein phosphorylation, through a signaling cascade that is activated by cellular dehydration. The in vitro data suggest the possibility of a different mode of activation—the enzyme is highly active in heterologous expression systems in assays 204 where the substrate MGDG is dispersed in detergent (Chapter 3). While an endogenous protein present in the heterologous host (yeast) used to express the SFR2/GGGT protein could provide the same activating PTM, this seems unlikely. Alternatively, the enzyme could be directly activated by mechanosensation of membrane destabilization. For example, when the GGGT/SF R2 protein is present in liquid crystal lamellar (bilayer) phase biomembranes, it is inactive. When conditions change, such as the formation of non-bilayer microdomains during membrane dehydration (e. g. during freezing), or detergent destabilization (e.g. the in vitro assay), a dynamic shift in equilibrium could result in the activation of GGGT/SFR2 to remodel MGDG into DGDG and oligogalactolipids. Examples of direct mechanosensation-based activation or inhibition of lipid metabolic enzymes, such as phospholipase A2, have long been known (Lehtonen and Kinnunen, 1995). While only circumstantial evidence currently exists for the possibility of GGGT/SFR2 to be activated through mechanosensation—this mode of activation is simpler and more direct than activation through PTMs—which may not be effective modes of regulation at sub-zero temperatures or states of severe dehydration during which GGGT/SFR2 is activated. This aspect of GGGT activity could be addressed by studying the activity in vitro as in Chapter 3, but where the enzyme has been reconstituted into large unilamellar vesicles (Szoka, Jr. and Papahadjopoulos, 1980). A preliminary experiment indicated that GGGT/SFR2 is inactive under these conditions (data not shown). With GGGT/SFR2 starting out inactive on in vitro vesicle bilayers, small molecules known to induce non-bilayer structure formation in vesicles could be added and the change in GGGT activity observed. Point or truncation mutants of the heterologous enzyme could be employed in identifying a potential mechanosensing 205 domain of GGGT, if initial results indicate its presence. Indeed, many other questions about GGGT enzymology could shed light on its overall function in plants. For example, GGGT/SFR2 is a glycosyl hydrolase, which has evolved high transglycosidase activity. Mutagenesis of the protein, followed by the standard activity assay could reveal residues or domains which are required for the high transglycosidasezhydrolase ratio exhibited by this enzyme. Results from both continued genetic study of SFR2 in planta and detailed enzymology of GGGT/SF R2 will provide great insight into the mechanisms through which membranes are actively remodeled in order to cope with changing environmental conditions. Triacylglycerol accumulation in Chlarnydomonas reinhardtii: a role for MLDP in lipid droplet biogenisis? The accumulation of triacylglycerols (TAGS) and lipid droplet composition in the green alga Chlamydomonas reinhardtii was investigated as described in Chapter 5. TAGS were found to accmnulate substantially during nitrogen deprivation (to ~65 % of total cellular fatty acids at 3 days -N), and accumulated in lipid droplets (LDs) within the cell. An LD fraction was purified, and the associated proteins identified through a mass spectrometric approach. An abundant LD protein, designated Major Lipid Droplet Protein (MLDP) was selected for further study. MLDP encodes a ~28 kDA protein with no known functional domains, and the MLDP mRNA was found to be strongly up- regulated during nitrogen deprivation. RNAi-mediated knockdown of MLDP resulted in a slight, but statistically significant increase in the average apparent lipid droplet diameter when compared to empty vector controls. 206 While this indicates a possible role for MLDP in lipid droplet morphology in Chlarnydomonas, this phenotype has been observed by knockdown of several different proteins in Drosophila which fall into broad functional classes (Beller e101,, 2008, Guo et al., 2008). As such, this phenotype on its own yields little mechanistic insight into the function of MLDP. As such, much work remains to be done on MLDP. Numerous attempts to over-express the MLDP protein as a GFP transgenic fusion failed—a common drawback of working in Chlarnydomonas, where successful transgene over- expression remains a community-wide bottleneck (Neupeit etal., 2009). As such, the confirmation of MLDP as an LD-associated protein will have to be conducted through more classical approaches, such as raising a specific anti-MLDP antibody for use in histochemical microscopy-based localization of MLDP, or through immuno-enrichment of the MLDP signal in LD fractions observed by western blot. Furthermore, potential functions for MLDP might be discovered by trying to complement mutants defective in various LD-associated proteins in other species such as yeast (Athenstaedt et al. , 1999), plants (Siloto et al., 2006), or fruit flies (Guo et al., 2008). Lastly, the potential for using homologous recombination in targeting MDP for insertional mutagenesis, though challenging, may be possible (Zorin et al., 2009). A true null allele of MLDP could provide greater insight into MLDP function, as the MLDP-RNAi knockdown level achieved was only ~60 % (Chapter 5). In addition to MLDP, many other factors are likely to be involved in TAG accumulation and LD formation during nitrogen deprivation in green algae. Due to the above-described limitations in reverse genetic approaches in Chlarnydomonas, development of forward genetic screens for both chemical and insertion mutants that are 207 defective in TAG accumulation or LD morphology will likely play a key role in the elucidation of the required factors for these proceses. The work described in this dissertation adds to the wealth of knowledge generated on plant lipid metabolism and how it is regulated during environmental stress. Furthermore, it provides insight into lipid responses during stress in green algae, an understanding of which is likely to play a key role in the development of algal species as feedstocks for renewable bioenergy. 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