REGULATION OF GLYCEROLIPID METABOLISM IN ARABIDOPSIS THALIANA By Anastasiya Lavell A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Biochemistry and Molecular Biology – Doctor of Philosophy 2019 REGULATION OF GLYCEROLIPID METABOLISM IN ARABIDOPSIS THALIANA ABSTRACT By Anastasiya Lavell Chloroplast membranes house the photosynthetic machinery and have a distinct lipid composition, with characteristically abundant galactolipids mono- and digalactosyldiacylglycerol (MGDG and DGDG). Both galactolipids are synthesized through both plastid and ER pathways in Arabidopsis, resulting in distinguishable molecular species. Phosphatidic acid (PA) is the first key intermediate formed by the plastid galactolipid biosynthetic pathway. It is further dephosphorylated to diacylglycerol (DAG), which is a substrate for MGDG Synthase (MGD1). MGD1 further adds a galactose to DAG from UDP-Gal. The enzymatic reactions yielding these galactolipids are known, but regulatory factors controlling this process are largely unknown. We identified a predicted rhomboid-like protease 10 (RBL10), located in plastids of Arabidopsis thaliana, that affects galactolipid biosynthesis likely through intramembrane proteolysis. Plants with T-DNA disruptions in RBL10 have greatly decreased 16:3 (acyl carbons : double bonds) and increased 18:3 acyl chain abundance in MGDG of leaves. Additionally, rbl10-1 mutant chloroplasts show reduced [14C]–acetate incorporation into MGDG during pulse- chase labeling, indicating a reduced flux through the plastid galactolipid biosynthesis pathway. While plastid MGDG biosynthesis is blocked in rbl10-1 mutants, they are capable of synthesizing PA, as well as producing normal amounts of total MGDG by compensating with ER-derived lipid precursors. These findings link this predicted protease to the utilization or transport of PA for plastid galactolipid biosynthesis potentially revealing a regulatory mechanism for galactolipid biosynthesis in chloroplasts. In addition to serving as a key metabolite, PA also has signaling roles in the cell, making its trafficking important to understanding plant cell metabolism. The substrate(s) of RBL10 are not yet known, but Blue-Native PAGE and FPLC analysis showed that RBL10 is part of a large molecular weight complex (>660kDa). The protein interactors of RBL10 are currently being probed by using co-immunoprecipitation and split-ubiquitin yeast two-hybrid approaches. Additionally, RBL10 seems to be autoproteolytic toward its own carboxy- terminal domain (CTD). The role of complex residency and autolytic activity of RBL10 is not currently clear, but these findings could help uncover the identity of a new transporter of PA in the chloroplast as well as a regulatory mechanism of its activity. ACKNOWLEDGEMENTS The journey to becoming a scientist was not trivial or quick, however, I could not see it happening any other way. Through the challenges of doctoral research, coursework, and the more bureaucratic aspects of belonging to a branch of science, I have gained an appreciation for the institution of academic research. On my way, I have encountered many individuals who deserve a sincere thank you, for their help, support, and guidance. I thank my advisor, Christoph Benning, who has given me not only the training to become a skilled researcher but also the room to grow and develop my own ideas. I always felt that Christoph truly cares for his students, and for that I am forever grateful. An unexpected way that Christoph helped me, was through having a lab full of great people. I want to thank all Benning lab members and I have to say a special thanks to Patrick Horn, Anna Hurlock, Yang Yang, and Kun Wang for answering a million questions, discussing my project endlessly, and making lab life fun. I want to recognize and thank my committee members, Jian Hu, Gregg Howe, Lee Kroos, and Danny Schnell. They have been insightful, constructive, and supportive throughout my career as a graduate student. I have immense gratitude for being a part of the Biochemistry and Molecular Biology department, and being supported by wonderful individuals like Jessica Lawrence, Ashley Parks, and Billy Yang. I have had the pleasure of working with Jon Kaguni and David Arnosti as graduate student directors and the honor of serving on the BMB Graduate Student Council. Doug Whitten has been instrumental for my doctoral work. In my future, I am not certain that I will ever find a collective of scientists as amazing as I did at the MSU-DOE Plant Research Laboratory. The PRL has been my home for the last iv four years and I will think fondly to my time here. I would not have been able to accomplish my work without the help of John Froehlich. I want to thank the PRL office staff for all their help and making my graduate experience a smooth one. I owe an immense thank you to all the undergraduate and high school students I have had the distinct pleasure of working with. In the hopes of teaching them I seemed to learn the most. For their mentorship and inspiration to teach, I thank Joyce Parker and Jon Stoltzfus. The Plant Biotechnology for Health Sustainability Training program greatly enriched my graduate life in numerous ways, and I have to thank Rob Last for making all that possible for me and many others. I cannot express my gratitude enough to my family and loved ones. The love and support from my family has been instrumental for my success. To my best friend and partner in life, Jonathan, I owe a huge thank you, I could not have done this without him. Last but not least, I want to thank my friends, those who have not wavered in their support of me. No challenge is too great when you have the love of your friends and family as well as the guidance and support from your mentors. v TABLE OF CONTENTS LIST OF TABLES ............................................................................................................ ix LIST OF FIGURES ........................................................................................................... x KEY TO ABBREVIATIONS ............................................................................................. xii CHAPTER 1 Rhomboids: Regulation from the Membrane .............................................. 1 Introduction ................................................................................................................. 2 iRhoms Play Regulatory Roles Through Non-Proteolytic Mechanisms ...................... 3 Prokaryotic Rhomboids ............................................................................................... 4 Eukaryotic Rhomboids ................................................................................................ 5 Modulating the Activity of Rhomboids ....................................................................... 10 REFERENCES ............................................................................................................... 13 CHAPTER 2 Cellular Organization and Regulation of Plant Glycerolipid Metabolism ... 19 Abstract ..................................................................................................................... 20 Introduction ............................................................................................................... 20 Lipid Transport Mechanisms ..................................................................................... 23 Regulation of Lipid Metabolism ................................................................................. 27 Lipid Signaling and Response to Stress ................................................................... 30 Spatial Arrangement and Complexes of Lipid Biosynthetic Enzymes ...................... 32 APPENDIX ..................................................................................................................... 35 REFERENCES ............................................................................................................... 38 CHAPTER 3 A Predicted Plastid Rhomboid Protease Involved in Phosphatidic Acid Metabolism in Arabidopsis thaliana ................................................................................ 48 Abstract ..................................................................................................................... 49 Introduction ............................................................................................................... 50 Results and Discussion ............................................................................................ 53 Discovery of a Role for RBL10 in Chloroplast Lipid Metabolism ......................... 53 JA and JA-Ile Levels are Affected in rbl10 .......................................................... 54 RBL10 is Located in the Inner Envelope Membrane of Chloroplasts .................. 55 ER-Assembled Lipids Serve as Primary Precursors for MGDG Biosynthesis in the rbl10-1 Mutant ............................................................................................... 56 PA Synthesized in Chloroplasts is Used for PG Biosynthesis but not MGDG .... 57 MGD1 Activity is not Affected by Loss of RBL10 ................................................ 58 PA Plastid Phosphatase Activity is Increased in rbl10-1 Mutants ....................... 58 A Refined Model of PA Metabolism in Arabidopsis Chloroplasts ........................ 59 Materials and Methods ............................................................................................. 61 T-DNA Lines ........................................................................................................ 61 Plant Growth Conditions ...................................................................................... 61 DNA Constructs and Complementation .............................................................. 62 vi Homology Modeling ............................................................................................. 62 Intact Chloroplast Isolation .................................................................................. 63 In Vitro Translation of Precursor Proteins ........................................................... 64 Import and Protease Protection Assays .............................................................. 64 Fractionation of Imported RBL10 ........................................................................ 65 Lipid Analysis ...................................................................................................... 65 Radiolabeling ....................................................................................................... 66 Acknowledgement .................................................................................................... 67 APPENDICES ................................................................................................................ 68 Appendix 3A Main Text Figures ................................................................................ 69 Appendix 3B Supplementary Figures ....................................................................... 76 REFERENCES ............................................................................................................... 81 CHAPTER 4 RBL10 Protein Properties and Oligomeric State ....................................... 87 Abstract ..................................................................................................................... 88 Introduction ............................................................................................................... 88 Results and Discussion ............................................................................................ 90 The Oligomeric State of RBL10 Indicates Stable Protein Interactors .................. 90 Co-Immunoprecipitation of RBL10 Yields Putative Protein Interactors ............... 91 RBL10 is Autolytic Towards its Carboxy-Terminal Domain ................................. 93 Catalytic Mutant Constructs of RBL10 Cannot be Stably Transformed into the rbl10-1 Mutant Background ................................................................................. 95 Materials and Methods ............................................................................................. 97 Plant Growing Conditions and Chloroplast Isolation ........................................... 97 SDS-PAGE and Immunoblot Analysis ................................................................. 97 Blue Native PAGE ............................................................................................... 98 Size Exclusion Chromatography ....................................................................... 100 Co-Immunoprecipitation .................................................................................... 101 Protein Production in E. coli .............................................................................. 103 Acknowledgement .................................................................................................. 105 APPENDIX ................................................................................................................... 106 REFERENCES ............................................................................................................. 112 CHAPTER 5 Arabidopsis DGD1 SUPPRESSOR 1 is a Subunit of the Mitochondrial Contact Site and Cristae Organizing System and Affects Mitochondrial Biogenesis ... 116 Abstract ................................................................................................................... 117 Introduction ............................................................................................................. 117 Results .................................................................................................................... 121 DGS1 is present in a multi-subunit protein complex with MIC60, TOM40, TOM20s and RISP ............................................................................................ 121 The dgs1-1 mutation alters the multi-subunit complex ..................................... 122 The dgs1-1 mutation alters mitochondrial size .................................................. 126 The dgs1-1 mutation alters mitochondrial lipid composition .............................. 128 The dgs1-1 mutation affects mitochondrial protein abundance, protein import and alternative respiratory capacity .......................................................................... 130 Higher drought stress tolerance in dgs1-1 mutant plant lines ........................... 132 vii Phylogenetic and co-expression networks of DGS1 putative orthologs ............ 136 Discussion ............................................................................................................... 137 Materials and Methods ........................................................................................... 142 Plant Materials ................................................................................................... 142 Plant Growth and Treatments ........................................................................... 143 Isolation of mitochondria, chloroplast and ER ................................................... 143 Blue-native PAGE ............................................................................................. 144 Immunoprecipitation (IP) ................................................................................... 144 Cross-Link Assay ............................................................................................... 144 Generation of the DGS1 antibody ..................................................................... 145 Immunoblot Analysis ......................................................................................... 146 Outer membrane ruptured mitochondria preparation and protease treatment .. 146 Protoplast Preparation and Fluorescence Microscopy ...................................... 147 Electron Microscopy .......................................................................................... 147 Lipid Extractions and Analysis ........................................................................... 148 Measurements of Respiratory Parameters on Plant Mitochondria .................... 148 In Vitro Mitochondrial Protein Import ................................................................. 149 In Situ Detection of Reactive Oxidative Species (ROS) .................................... 150 Dark-induced Senescence ................................................................................ 150 Quantification of H2O2 ....................................................................................... 150 Relative Water Content Analysis ....................................................................... 151 Chlorophyll analysis and fluorescence .............................................................. 151 Gas Exchange ................................................................................................... 152 Total RNA Preparation and cDNA Synthesis .................................................... 152 Global Transcript Analysis ................................................................................. 152 Mass Spectrometry ............................................................................................ 155 Phylogenetic analysis ........................................................................................ 155 Co-expression analysis ..................................................................................... 156 Accession Numbers........................................................................................... 156 Supplemental Data Sets .................................................................................... 156 Acknowledgement .................................................................................................. 157 Author Contribution ................................................................................................. 158 APPENDICES .............................................................................................................. 159 Appendix 5A Main Text Figures ........................................................................ 160 Appendix 5B Supplemental Figures .................................................................. 175 REFERENCES ............................................................................................................. 182 CHAPTER 6 Future Perspectives ................................................................................ 190 Introduction ............................................................................................................. 191 Role of Proteolysis in Lipid Homeostasis ................................................................ 191 RBL10 and its Protein Interactome ......................................................................... 193 Role for Autocatalysis of RBL10 ............................................................................. 195 Responses to Stress and Lipid Remodeling ........................................................... 197 Concluding Remarks .............................................................................................. 198 REFERENCES ............................................................................................................. 199 viii LIST OF TABLES TABLE 4.1. Co-Immunoprecipitation with RBL10-YFP-HA using isolated chloroplast protein lysate, results in recurring appearance of select proteins ................................ 107 ix LIST OF FIGURES Figure 2.1. Scheme of phosphatidic acid (PA) synthesis in the plastid and the endoplasmic reticulum (ER) ........................................................................................... 36 Figure 2.2. Proposed movement of galactolipids after synthesis from diacylglycerol (DAG) ............................................................................................................................. 37 Figure 3.1. Structure and phenotypes of rbl10 mutant alleles ........................................ 70 Figure 3.2. Localization of RBL10 to the Inner Envelope Membrane of Chloroplast ..... 71 Figure 3.3. Radiolabeled pulse chase of detached leaves using 14C-acetic acid ........... 72 Figure 3.4. Labeling of isolated chloroplasts of rbl10-1 mutant and WT with 14C-acetate . ....................................................................................................................................... 73 Figure 3.5. Probing PA metabolism in rbl10-1 ............................................................... 74 Figure 3.6. Proposed model of PA metabolism in the chloroplast envelope membranes .. ....................................................................................................................................... 75 Figure 3.S1. Full lipid analysis of wild-type (WT) and rbl10 leaves ................................ 77 Figure 3.S2. Acyl composition of rbl11 total leaf lipids and MGDG ................................ 78 Figure 3.S3. Changes in oxylipin levels in the rbl10 mutants ......................................... 79 Figure 3.S4. Seed phenotypes of rbl10 plants ............................................................... 80 Figure 4.1. Blue Native PAGE of solubilized chloroplasts from WT, rbl10-1 mutant, and rbl10-1 complemented lines producing RBL10-YFP-HA protein .................................. 108 Figure 4.2. Size Exclusion Chromatography of protein standards and solubilized chloroplasts of rbl10-1:RBL10-YFP-HA line show that RBL10 migrates in a complex at a high MW ....................................................................................................................... 109 Figure 4.3. Production of C-Terminally tagged RBL10-T in E. coli .............................. 110 Figure 4.4. The fusion protein of MBP-RBL10 WT and single catalytic mutant can be detected with anti-MBP antibody .................................................................................. 111 x Figure 5.1. DGS1 is present in a large multi-subunit protein complex with MIC60, TOM40, TOM20s and RISP ......................................................................................... 161 Figure 5.2. A single point mutation in DGS1 alters the multi-subunit complex ............ 162 Figure 5.3. The dgs1-1 mutation alters mitochondrial size .......................................... 164 Figure 5.4. The dgs1-1 mutation alters mitochondrial lipid composition ...................... 166 Figure 5.5. The dgs1-1 mutation affects mitochondrial protein abundance, protein import and alternative respiratory capacity ............................................................................. 168 Figure 5.6. The dgs1-1 mutation imparts higher drought stress tolerance ................... 170 Figure 5.7. Transcriptome analysis of dgs1 mutants ................................................... 171 Figure 5.8. Phylogenetic analysis of DGS1- and NCA2-related proteins ..................... 173 Figure 5.S1. The intramitochondrial localization of TOM20-2 and TOM40 in Col-0 and dgs1 mutant lines ......................................................................................................... 176 Figure 5.S2. Related to Figure 4A and 4B. Leaf lipid analysis of wild type and mutant lines .............................................................................................................................. 177 Figure 5.S3. The defective growth phenotype of mutants producing the DGS1-1 protein . ..................................................................................................................................... 179 Figure 5.S4. Analyses of hydrogen peroxide in dgs1-1 lines ....................................... 181 xi KEY TO ABBREVIATIONS ATS1 – glycerol-3-phosphate acyltransferase DAG – diacylglycerol DGD1 – digalactosyldiacylglycerol synthase 1 DGDG - digalactosyldiacylglycerol DGS1 – suppressor of DGD1 ER – endoplasmic reticulum FA – fatty acid GlpG – E. coli rhomboid JA – jasmonic acid JA-Ile – jasmonic acid isoleucine MGD1 – monogalactosyldiacylglycerol synthase 1 MGDG – monogalactosyldiacylglycerol PC – phosphatidylcholine RBL10 – rhomboid-like protein 10 TAG – triacylglycerol PA – phosphatidic acid PARL – presenilin-associated rhomboid-like protein PG – phosphatidylglycerol TeGDG – tertragalactosyldiacylglycerol TGD1 – TRIGALACTOSYLDIACYLGLYCEROL 1 TGDG – trigalactosyldiacylglycerol xii CHAPTER 1 Rhomboids: Regulation from the Membrane 1 Introduction Found in nearly all organisms, a superfamily of rhomboid proteases carries out regulated intramembrane proteolysis. These serine-type endopeptidases utilize a serine-histidine catalytic dyad to cleave transmembrane segments of peptides in the hydrophobic environment of the membrane. The founding member of this family was originally discovered in Drosophila melanogaster where it cleaves the factor Spitz from the Golgi membrane and initiates the epidermal growth factor signaling pathway (Lee et al., 2001, Urban et al., 2001, Urban et al., 2002). Since the discovery of this family, rhomboid-like proteins have been predicted across nearly all organisms including animals, plants, bacteria, protozoa, and fungi. With more available genomic data, subclasses of rhomboid or rhomboid-like proteins were identified, creating subtypes with and without the catalytic residues as well as varied amino and carboxy-terminal sequences (Lemberg and Freeman, 2007). Although one of the first characterized rhomboids was in D. melanogaster, the origin of the superfamily is hypothesized to date back to a bacterial ancestor with multiple horizontal gene transfer events giving rise to the RHO subfamily and then separately the Presenilin-associated rhomboid-like protein (PARL) subfamily (Koonin et al., 2003). Major strides have been made in the characterization of rhomboid family proteins in diverse organisms, yet with so many predicted rhomboid encoding genes yet to be investigated, a lot of opportunities remain for new and unique regulatory examples. This chapter presents examples of rhomboids and their biological roles from representative organisms across kingdoms and highlights the gaps in knowledge on rhomboid function in plants. 2 iRhoms Play Regulatory Roles Through Non-Proteolytic Mechanisms There are two main types of rhomboid-like proteins, active proteases and inactive iRhoms. Though inactive, iRhom proteins possess many of the characteristics of active rhomboids and display the rhomboid transmembrane architecture. However, they have lost the conserved serine and histidine residues which confer proteolytic activity. (Lemberg and Freeman, 2007). Despite a lack of enzymatic activity, there is growing evidence that iRhoms play important roles in metabolism. In flies and mammals, iRhoms seem to participate in ER-associated degradation of EGF and EGF-like ligands, contrary to the effects of active rhomboid-1 on promoting the EFG signaling pathway in Drosophila (Zettl et al., 2011). In mice, the roles of iRhom1 and 2 were shown to determine the trafficking of a shedding protease TACE/ADAM17 in macrophages, where proper iRhom mediated trafficking through the ER results in the processing and release of tumor necrosis factor (TNF) (Christova et al., 2013). Though the relationship between iRhom1, iRhom 2, and TACE is well supported, knock-out (KO) mice of iRhom1 fail to create white fat deposits and die (Christova et al., 2013), where iRhom2 KO mice appear to exhibit inflammatory defects only (Adrain et al., 2012). These varied phenotypes of homozygous KOs of iRhom1 and 2, suggest possible differences in expression or additional functions beyond TACE trafficking. The effects on lipid deposition in mice was further investigated in relation to iRhom2, where possible therapeutic strategies for myocardial inflammation resulting from obesity were found by inhibiting endoplasmic reticulum stress (ERS) and iRhom2 signaling pathways (Ge et al., 2019). With a large number of iRhoms predicted alongside active rhomboids, their biological roles remain an open field of research. 3 Prokaryotic Rhomboids Bacteria - In the microorganism, Providencia stuartii, the rhomboid AarA has been shown to play a role in the oligomerization of the twin-arginine translocase complex. The complex is made up of several Tat proteins, of which one serves as the recruiting factor, TatA. This small peptide has a single pass transmembrane helix which is a substrate for AarA. Upon cleavage, the processed TatA recruits the other Tat proteins and the twin-arginine translocase complex is oligomerized (Stevenson et al., 2007). The AarA gene was first discovered through a mutagenesis screen searching for regulatory components of aminoglycoside acetyltransferase, which is involved in resistance to the respective broad- spectrum antibiotic. The resultant aarA mutants had increased resistance to gentamicin as well as changes in cell morphology (Rather and Orosz, 1994). Contrary to the AarA, the natural substrate of the Escherichia coli rhomboid GlpG and its endogenous role has not yet been identified. Though bacteria lacking GlpG don’t share all the phenotypes as the aarA mutants, introducing the GlpG gene rescues the aarA mutants (Clemmer et al., 2006). Despite our lack of understanding of its biological function, the numerous crystal structures, models, (Wang et al., 2006, Wang et al., 2007, Ha et al., 2013, Vinothkumar et al., 2013) and kinetic assays of GlpG (Maegawa et al., 2005, Xue and Ha, 2012, Cho et al., 2016, Guo et al., 2016), led to a better understanding of substrate recognition mechanisms of rhomboid proteases as well as substrate cleavage. Another bacterial rhomboid, GluP in Bacillus subtilis, has been implicated in glucose transport as well as cell division. However, the mechanism of action and its natural substrate remains to be discovered (Mesak et al., 2004). 4 The ability to study the kinetics and structure of GlpG generated a lot of knowledge about the catalytic mechanisms of rhomboids. However, with the emerging diversity of these proteases in eukaryotic organisms a diversity in molecular action might be expected as well. Eukaryotic Rhomboids Animals – Following the discovery of the Drosophila rhomboid-1, evidence became available that a human rhomboid RHBDL2 is capable of activating EGF ligands and EGFR signaling, though, since there are other parallel pathways to EGF activation, the significance of this finding still remains to be clarified (Adrain et al., 2011). In mitochondria of human neuron cells, a PARL type rhomboid performs an important role in determining the trafficking of a factor called PINK1. The processing of the single transmembrane domain of PINK1 can alter its subcellular location, where the non-cleaved form is targeted to the outer membrane of the mitochondrion and the cleaved form of Pink1 is targeted to the inner envelope membrane of the mitochondrion (Meissner et al., 2011). In the outer membrane location, Pink1 recruits Parkin, a ubiquitin ligase, and initiates mitophagy. In this instance, unhealthy mitochondria are turned over through Pink1/Parkin directed mitophagy. When Pink1 is cleaved and targeted to the inner membrane, it is ultimately degraded by the 26S proteasome. The determining factor for PARL cleavage is the phosphorylation status of the amino-terminal domain (NTD) of PARL. Under normal ATP levels in the mitochondrion matrix, a kinase, PDK2, will phosphorylate PARL, thereby preventing PARL from self-processing to a less active form PACT (Jeyaraju et al., 2006). Under reduced ATP conditions, PARL will not be phosphorylated to the same extent and 5 will auto-catalytically remove its own NTD, resulting in reduced Pink1 cleavage and ultimately initiation of mitophagy (Meissner et al., 2015, Shi and McQuibban, 2017). The autolytic activity of PARL (b-cleavage) occurs after the initial removal of mitochondrial targeting peptide (a-cleavage) and generates a peptide that is targeted to the nucleus (Sik et al., 2004). This knowledge on proteolytic control of PINK1 trafficking and its influence on initiation of mitophagy provides insight on the key roles of rhomboid-like proteins in complex biological pathways. The change in phosphorylation status of PARL and the subsequent autolytic b-cleavage to the less active form PACT is one example of the self-regulation of rhomboid function. More importantly, these studies demonstrate a mechanism by which rhomboids may act as transducers of metabolic status in the cell or organelle. Protozoa – The genome of the apicomplexan Plasmodium falciparum, implicated in the human disease malaria, also encodes active rhomboid proteases. Proteases PfROM1 and PfROM4 exhibit sheddase activity and cleave a diverse array of adhesin proteins present on the surface of the pathogen; this shedding of adhesins is important during completion of host cell invasion and membrane fusion (Baker et al., 2006). Furthermore, PfROM4 exhibits high specificity for particular adhesins such as EBA-175. Performing mutagenesis of the adhesin protein leads to an inhibition of cleavage by PfROM4 and thereby compromises host cell invasion (O'Donnell et al., 2006). In similar fashion, the rhomboids of Toxoplasma gondii are important for the shedding of adhesins during invasion of host cells. Five rhomboid-like proteins and a single PARL-type protein were predicted in T. gondii, where proteolytically active TgROM5 appeared to be a spatially and temporally fitting candidate to aid in invasion (Brossier et al., 2005). Further 6 work showed that TgROM4 also aids in invasion by processing adhesins on the cell surface of T. gondii (Buguliskis et al., 2010). Rhomboids in other protozoans have been predicted and understanding the role of rhomboids in host cell invasion creates exciting opportunities for novel drug targets in cases of disease-implicated protozoa such as Plasmodium. Fungi - A pathogenic mold, Aspergillus fumigatus, is responsible for over 90% of pulmonary mycosis and can colonize the lung tissues of immunocompromised humans through not yet well established molecular mechanisms (Latge, 2001). During colonization of lung tissue, Aspergillus must adapt to lower levels of oxygen than that of the atmosphere. A key protein in the Aspergillus genome, identified in mediating this adaptation, is a sterol regulatory element-binding protein (SREBP) homolog SrbA, without which the pathogen was unable to grow under hypoxic (Willger et al., 2008). It was later identified that SrbA is likely a substrate of an Aspergillus rhomboid protease RbdA. Under hypoxic conditions, the rbdA mutant can be rescued by overexproduction of a truncated N-terminal fragment of SrbA in the nucleus (Vaknin et al., 2016). The regulation of adaptability and virulence of a pathogenic mold that is in part controlled by a rhomboid protease, creates yet another potential drug target, in this case for treating immunocompromised individuals infected with Aspergillus. Work in fission yeast described a rhomboid Rbd2 which interacts with the AAA- ATPase Cdc48 in order to identify its ubiquitinated substrate, a SREBP. The Cdc48/Rbd2 interaction is important in the processing of SREBPs and disrupting the binding of Cdc48 to the C-terminus of Rbd2 results in degradation of the ubiquitinated SREBP (Hwang et al., 2016). In both fission yeast and mold, there seem to be additional, diverse terminal 7 domains that mediate protein-rhomboid interactions. The necessity of Cdc48 for substrate identification by Rbd2 is uniquely different from the presumed rhomboid-substrate interaction within the membrane by virtue of “scanning” the membrane. Though it has been proposed that scanning by a rhomboid in the membrane may be occurring at biologically relevant rates (Kreutzberger et al., 2019), with the diversity of rhomboid mechanisms found so far, it’s likely that more rhomboid chaperones will soon emerge. Plants - Though an appreciable number of rhomboid mechanisms have been elucidated, there is yet to be a rhomboid and endogenous substrate pair identified in plants. Numerous members of the family have been predicted in model species Arabidopsis thaliana, Populus trichocarpa, and in Oryza sativa (Garcia-Lorenzo et al., 2006). In Arabidopsis, 20 rhomboid-encoding genes were predicted and 18 in rice (Tripathi and Sowdhamini, 2006). Though all predicted genes encoded proteins displaying the rhomboid fold, the number of active rhomboids containing the conserved catalytic domains has been refined to 13 in Arabidopsis and 12 in rice (Lemberg and Freeman, 2007) with the remaining predicted genes to produce non-catalytic members of the family. The subcellular distribution of rhomboids in plants has been speculated for some members and demonstrated experimentally for others. Since their function is tied to their membrane location, at least for the process of proteolysis, ascertaining their subcellular distribution is highly important for identifying endogenous substrates. Of the 13 predicted rhomboids in Arabidopsis, two are shown to be Golgi-associated (Kanaoka et al., 2005), two are associated with the chloroplast (Knopf et al., 2012, Thompson et al., 2012), one mitochondrial PARL member is confirmed (Knopf et al., 2012), two rhomboids are 8 predicted at the plasma membrane (Benschop et al., 2007, Inze et al., 2012), and the others have mixed predictions and lack experimental evidence to support their locations at this time. So far, only one rhomboid in plants has been shown to possess regulated intermembrane proteolysis. The Golgi localized AtRBL2 can process the Drosophila RHO-1 substrates Spitz and Keren; this finding was in contrast to another Golgi resident, AtRBL1, which could not process either Spitz or Keren despite its sequence similarity to DmRHO-1 (Kanaoka et al., 2005). Though the catalytic activity of these two rhomboids has been shown, there are no current hypotheses on what their endogenous substrates are nor what their overall physiological role is. However, this analysis does confirm that rhomboid-like proteins can in fact carry out intramembrane proteolysis in plants. For the biological importance of plant rhomboids, the best studied member is RBL10 (At1g25290). Knock-out mutants of RBL10 were first characterized for their developmental defects, showing irregular flower formation, abnormal pollen morphology, and reduced number of seeds per silique formed (Thompson et al., 2012). The overall mechanism of how RBL10 influenced these developmental processes was not further elucidated. However, there was a hypothesized disruption of the plant hormone jasmonic acid (JA) signaling in this mutant, which could potentially explain the phenotypes. Further evidence was provided through a proteomic approach, where levels of allene oxide synthase (AOS), an enzyme catalyzing a key step in JA production, were reduced in rbl10/rbl11 double mutants compared to wild type (Knopf et al., 2012). However, due to the use of a double knock-out mutant for the proteomic analysis, it was unclear which rhomboid actually influenced the levels of AOS. Though redundancy is possible, it was 9 recently shown that RBL10 participates in lipid homeostasis of Arabidopsis while RBL11 does not. Knock-out mutants of rbl10 show an impaired ability to utilize chloroplast synthesized phosphatidic acid (PA), a key intermediate in lipid biosynthesis, for the formation of galactolipids such as monogalactosyldiacylglycerol (MGDG) (Lavell et al., 2019) (see Chapter 2). This led to the hypothesis that RBL10 may affect trafficking of PA across the inner envelope membrane of the chloroplast. Though the relationship between the reproductive phenotypes and lipid metabolism does not seem immediately clear, the bridge between the two may after all be JA. Since JA is synthesized from chloroplast lipid derived fatty acids, and rbl10 mutant shows a slightly attenuated JA response after wounding (Lavell et al., 2019), it is possible that reduced JA supply during the dynamic process of flower and pollen tube expansion could lead to the developmental phenotypes previously observed. While many of the rhomboids studied in other organisms seem to play important roles in disease development, the rhomboids in plants are poised to potentially be of importance to plant growth and development. Modulating the Activity of Rhomboids As roles of rhomboids are uncovered in various human diseases, the ability to modulate their activity has become an important area of research. The regulation of rhomboid activity is varied; an interesting example is found in human cells where RHBDL4, an ER localized rhomboid-like protein, binds cholesterol and appears to be modulated in activity by the levels of cholesterol present (Paschkowsky et al., 2018). Aside from self-regulation 10 through autolytic activity as seen in the human PARL, rhomboid catalysis can be inhibited by a limited number of chemical protease inhibitors. Due to the hydrophobic nature of the active site of rhomboids, not all serine-type protease inhibitors are effective. An inhibitor class derived from natural rhomboid substrates, peptidyl-chloromethylketones (CMK) has been shown to inhibit GlpG through in vivo and in vitro assays (Zoll et al., 2014). A group of compounds called ß-lactams was identified to be effective in inhibiting AarA and GlpG. However, interestingly there was some enhanced inhibition of GlpG compared to AarA, raising the exciting possibility of selective inhibition of individual rhomboids (Pierrat et al., 2011). A related class of compounds called ß-lactones has also been classified as inhibitors of rhomboid activity (Wolf et al., 2015). Two more compounds, 3,4-dichloroisocoumarin (DCI) and diisopropyl fluorophosphonate (DFP), were shown to be effective in inhibiting GlpG in in vitro assays, with DFP being more potent and causing irreversible inhibition of GlpG (Xue and Ha, 2012). Despite the discovery of potential inhibitors of rhomboids, many of the compounds do not lead to complete ablation of activity and are reversible over time. Even for the more effective compounds, cross reactivity with other vital serine endopeptidases has not been well investigated and remains a hurdle to overcome. Aside from medical implications of rhomboid inhibitors, the discovery and use of inhibitors has provided details about the way rhomboids interact with substrate-like compounds, giving insights into their catalytic mechanisms. For a more detailed overview on rhomboid inhibitors, the reader is directed to an article by Kvido Strisovsky (Strisovsky, 2016). Overall, understanding the molecular action and biological roles of rhomboids could lead to novel targets for treatments of human disease. Furthermore, understanding 11 rhomboids in plants could help identify novel regulatory mechanisms for growth and development as well as stress response. These research areas will likely be important for human health and agricultural productivity. Further research on the involvement of rhomboids in lipid metabolism is needed to understand how they may impact plant oil accumulation as well as photosynthetic efficiency of Arabidopsis but ultimately how this may translate to other agriculturally valuable crops. RBL10 is a first step to uncovering novel regulatory mechanisms in the complex processes of PA and galactolipid metabolism. 12 REFERENCES 13 REFERENCES Adrain, C., Strisovsky, K., Zettl, M., Hu, L., Lemberg, M.K. and Freeman, M. (2011) Mammalian EGF receptor activation by the rhomboid protease RHBDL2. Embo Rep, 12, 421-427. Adrain, C., Zettl, M., Christova, Y., Taylor, N. and Freeman, M. (2012) Tumor necrosis factor signaling requires iRhom2 to promote trafficking and activation of TACE. Science, 335, 225-228. Baker, R.P., Wijetilaka, R. and Urban, S. (2006) Two Plasmodium rhomboid proteases preferentially cleave different adhesins implicated in all invasive stages of malaria. Plos Pathog, 2, 922-932. 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Embo J, 33, 2408-2421. 18 Cellular Organization and Regulation of Plant Glycerolipid Metabolism CHAPTER 2 This work has been further edited and is published in Plant and Cell Physiology (https://academic.oup.com/pcp) Lavell, A. and Benning, C. (2019) Cellular Organization and Regulation of Plant Glycerolipid Metabolism. Plant Cell Physiol, 60, 1176-1183. 19 Abstract Great strides have been made in understanding how membranes and lipid droplets are formed and maintained in land plants, yet much more is to be learned given the complexity of plant lipid metabolism. A complicating factor is the multi-organellar presence of biosynthetic enzymes and unique compositional requirements of different membrane systems. This necessitates a rich network of transporters and transport mechanisms that supply fatty acids, membrane lipids, and storage lipids to their final cellular destination. Though we know a large number of the biosynthetic enzymes involved in lipid biosynthesis and a few transport proteins, the regulatory mechanisms, in particular, coordinating expression and/or activity of the majority remain yet to be described. Plants undergoing stress alter their membranes’ compositions, and lipids such as phosphatidic acid have been implicated in stress signaling. Additionally, lipid metabolism in chloroplasts supplies precursors for jasmonic acid (JA) biosynthesis, and perturbations in lipid homeostasis has consequences on JA signaling. In this review, several aspects of plant lipid metabolism are discussed that are currently under investigation: cellular transport of lipids, regulation of lipid biosynthesis, roles of lipids in stress signaling, and lastly the structural and oligomeric states of lipid enzymes. Introduction Lipid metabolism in plants is an essential process that provides cells with membranes, a storage form of energy and building blocks, and potent signaling compounds. In this review, we will give some necessary background on plant lipid metabolism and outline the frontiers of our knowledge on plant lipid biosynthesis and transport. In addition, we 20 will point out the outstanding questions about the biophysical arrangement or topology of lipid biosynthetic enzymes. Plant cells, as any other eukaryotic cell, are composed of many membrane systems and compartments with unique lipid compositions. Here we will focus primarily on land plants and how they carry out lipid biosynthesis in the plastid and the endoplasmic reticulum (ER), and related processes associated with those compartments. Most of the information described here is based on the model plant Arabidopsis thaliana (Arabidopsis) unless otherwise indicated. In both the ER and plastid, phosphatidic acid (PA) is the first lipid species formed by de novo complete acylation of glycerol-3-phosphate (glycerol-3-P). During the biosynthesis of PA, the specificity of the acyltransferases associated with the respective compartments determines the length of acyl chains esterified. In the plastid, glycerol-3-P acyl transferase and lysophosphatidic acid acyltransferase (ATS1 and ATS2) produce a PA bearing 18:1 and 16:0 acyl chains (carbon # : double bond #) at the sn-1 and sn-2 positions of the glycerol backbone, respectively (Bin et al., 2004, Kim and Huang, 2004, Tzafrir et al., 2004, Xu et al., 2006). Meanwhile, the ER acyltransferases produce a PA with predominantly 18-carbon acyl chains at both sn-1 and sn-2 positions (Kim et al., 2005). This distinction results in two identifiable sets of molecular species of lipids that are, within limitations, representative of the lipid assembly in the respective compartments. Though the fluxes through these two parallel pathways have been estimated based on the 16/18 carbon ratio, or more precisely by positional analysis of membrane lipid acyl composition, there are some caveats on the specificity of acyltransferases and possible lipid reassembly. For example, acyl editing of chloroplast- assembled lipids has recently been observed through in vivo lipid tagging using an ER- 21 targeted D6-desaturase from Physcomitrella patens (Hurlock et al., 2018). Already, acyl editing of ER lipids has been known for some time for phosphatidylcholine (PC) in plants (Bates et al., 2007). However, if acyl editing of chloroplast lipids contributes significantly to the acyl composition of chloroplast lipids, the reliability of the 16/18 carbon ratio as a simple metric for the origin of plastid lipids needs to be reevaluated. This is particularly true for Chlamydomonas, for which a plastid-type lysophosphatidic acid acyltransferase associated with the ER has been discovered (Kim et al., 2018), explaining the unusually high 16/18 ratio of Chlamydomonas galactolipids, even though it has been shown to import ER-derived lipid precursors into the plastid (Warakanont et al., 2015). In addition to differing acyltransferase preferences of the two pathways, the conversion of PA to other lipid classes also differs between the two compartments as the ER produces exclusively phospholipids and triacylglycerols while the plastid synthesizes the phospholipid phosphatidylglycerol (PG), the sulfolipid sulfoquinovosyldiacylglycerol (SQDG), and the two galactolipids mono- and digalactosyldiacylglycerol (MGDG and DGDG). Though components of both pathways are present in most land plants, not all plants utilize them both or to the same extent. For example, grasses primarily rely on the ER pathway (Petroutsos et al., 2014, Yang et al., 2017). However, to understand the interplay between the ER and plastid pathways, Arabidopsis provides an excellent starting model due to its near equal utilization of both pathways. 22 Lipid Transport Mechanisms As pointed out above, Arabidopsis produces glycerolipids through the plastid and ER pathways, also denoted prokaryotic and eukaryotic pathways, respectively (Browse et al., 1986, Kunst et al., 1988, Mongrand et al., 1998). There is also some lipid assembly in the mitochondrion and the reader is directed to the Acyl-Lipid Metabolism chapter of the Arabidopsis Book (Li-Beisson et al., 2013) for further information concerning mitochondrial lipid metabolism. Though the assembly of glycerolipids occurs in more than one compartment, with the exception of fatty acids synthesized in the mitochondrion mostly as precursors to lipoic acid (Wada et al., 1997), the plastid is the location where the vast majority of fatty acids are de novo synthesized. In order to supply substrates to the glycerolipid assembly machinery at the ER, acyl groups must be exported from the plastid. In addition, the ER provides glycerolipid precursors to the chloroplast requiring a transport system. The flux of acyl groups from the plastid is substantial especially in seed oil storage tissues, and yet at this time it is still not unambiguously known by which molecular mechanism the acyl groups are exported from the plastid (Fig. 2.1) (Koo et al., 2004). One hypothesis suggests transport of lipids or fatty acids through ER-plastid contact sites, which have been observed (Andersson et al., 2007). An inner chloroplast envelope membrane-spanning protein FAX1 seems to play a role in lipid homeostasis of ER-derived lipids and can complement the fat1 yeast mutant deficient in fatty acid export (Li et al., 2015). Although FAX1 is likely to aid in the transport of fatty acids across the inner envelope membrane at least in specific tissues, it appears as though not all fatty acid export from chloroplasts is abolished in the fax1 mutant and other mechanisms may be in play. The necessity of transport channels or other protein-mediated mechanisms to 23 move fatty acids across the inner envelope membrane as well as the outer envelope leaflets to the ER membranes (Fig. 2.1) makes this an active area of research. After lipids are assembled at the ER, there are several proposed mechanisms on how those lipids make it back to the plastid. One reported transport mechanism involves the phospholipid transporter ALA10. Mutants of ALA10 have a lowered MGDG/PC ratio and ALA10 overexpression leads to an increase in MGDG/PC ratio, suggesting a role of ALA10 in maintaining MGDG metabolism in the chloroplast (Botella et al., 2016, Botella et al., 2017). Based on these data it was concluded that this transporter may be important for efficient translocation of PC across the ER membrane and to the plastid either through ER-plastid contact sites or by another mechanism yet to be described. A means of reimporting ER-assembled lipids into the plastid has been described and a multiprotein ABC transporter complex consisting of three proteins TRIGALACTOSYLDIACYLGLYCEROL1, 2, 3 (TGD1, 2, 3) has been identified. Together with two additional proteins, TGD4 and TGD5, these proteins provide a conduit for lipid transfer that spans the outer and inner envelope membranes of the plastid and coordinates lipid import from the ER in a unidirectional manner (Xu et al., 2003, Xu et al., 2005, Awai et al., 2006, Lu et al., 2007, Lu and Benning, 2009, Xu et al., 2010, Roston et al., 2011, Roston et al., 2012, Fan et al., 2015). TDG4 and TGD5 are located at the outer envelope, where TGD4 contains a PA binding site (Wang et al., 2013) and is hypothesized to facilitate the translocation of ER-assembled PA to the plastid membranes (Wang et al., 2012). This occurs in coordination with the inner envelope subcomplex composed of TGD1, TGD2, and TGD3, which may also assist the translocation of PA through the identified PA-binding site of the TGD2 protein (Awai et al., 2006, Lu and Benning, 2009). 24 Since PA is also synthesized at the chloroplast stroma side of the inner envelope membrane, there could be an outwards transporting mechanism that translocates plastid PA and inserts it into the same leaflet as the ER-sourced PA. A bacterial ABC transporter of lipopolysaccharide (LPS) precursors resembling the plant TGD complex has been shown to mediate the outwards transport of lipids from the inner membrane to the outer membrane of the cell (Sperandeo et al., 2008). Bacterial lipid transporter systems Mla, Pqi and Yeb, have been structurally analyzed and have intermembrane spanning projections, like the LPS transporter and the TGD complex, which aid in lipid transport from the inner to the outer membrane (Ekiert et al., 2017). It is possible that similar mechanisms exist to move plastid-assembled lipids outwards through the chloroplast to the envelope membranes (Fig. 2.1). As both eukaryotic and prokaryotic DAG species are galactosylated by monogalactosyldiacylglycerol synthase 1 (MGD1) located at the intermembrane leaflet of the inner envelope membrane, the transport of the product MGDG to the thylakoid membranes needs to be aided by a protein channel or transport mechanism. In addition to MGDG, the product of digalactosyldiacylglycerol synthase 1 (DGD1), digalactosyldiacylglycerol (DGDG), must also be translocated across the outer and inner envelope membranes to be incorporated into the thylakoid membranes. These galactolipid transport mechanisms could share the same machinery or use specific transporters for each lipid (Fig. 2.2). A proposed mechanism by which DGDG moves from the outer envelope to the inner envelope membrane involves the N-terminal extension of DGD1 itself (Kelly et al., 2016) (Fig. 2.2). 25 Another intriguing area of study is the export of plastid lipids under certain environmental stresses such as phosphate starvation. A study looking for a non-DGD1 dependent pathway of DGDG synthesis led to the discovery of a suppressor mutant dgs1 which accumulates galactolipids despite lacking the enzyme DGD1 responsible for the bulk synthesis of DGDG (Xu et al., 2008, Moellering and Benning, 2010). Interestingly, the homozygous dgd1/dgs1 mutant also accumulated oligogalactolipids (Xu et al., 2008) such as trigalactosyldiacylglycerol (TGDG) which are typically induced by freezing stress (Moellering et al., 2010). Under phosphate starvation, Arabidopsis plants substitute phospholipids for galactolipids in order to maintain cellular membranes while reserving phosphate for DNA and protein synthesis (Härtel et al., 2000, Härtel et al., 2001, Kobayashi et al., 2009). During early stages of phosphate starvation, PC abundance, unlike other phospholipids, increases and so does DAG that may be derived from the PC before glycolipid synthesis increases (Jouhet et al., 2003). There is some evidence that a mitochondrial complex containing Mic60 plays a role in the response to phosphate starvation and import of plastid-derived galactolipids to the mitochondrion under phosphate starvation (Michaud et al., 2016). It is also hypothesized that MGDG is important during pollen tube expansion to satisfy the need for lipids for rapidly growing membranes (Kobayashi et al., 2004). Galactolipids like DGDG can be detected by antibody in the plasma membrane and the abundance of MGD synthases 2 and 3 is high in pollen tubes. The use of a synthetic MGD synthase inhibitor galvestine-1 showed reduced pollen tube elongation (Botte et al., 2011). These are a few examples of the presence of galactolipids in extraplastidic membranes requiring export from the plastid. 26 Presumably, the ER as well as other cellular membranes would acquire galactolipids by mechanisms similar to those apparent during phosphate-deprived conditions. Regulation of Lipid Metabolism For an integral process such as membrane biogenesis, there must be tight regulation of the flux through the lipid biosynthetic pathways. However, surprisingly little is known about the means by which lipid-synthesizing enzymes are regulated with regard to their genes’ expression or their enzymatic activity. One of the better understood regulatory mechanism of lipid metabolism is the regulation of the heteromeric acetyl-CoA carboxylase (ACCase), the complex initiating fatty acid synthesis within plastids. The expression of ACCase subunit genes is coordinated and it appears that the amount of each subunit is modified post-translationally as subunits oligomerize and excess proteins are degraded (Sasaki and Nagano, 2004) and the protein activity is stimulated by light (Sasaki et al., 1997). Like in animals and yeast, plant ACCase enzymes in Arabidopsis, Nicotiana, as well as Brassica, have been shown to catalyze the committed step of fatty acid synthesis, and respond to feedback inhibition by fatty acids (Shintani and Ohlrogge, 1995, Andre et al., 2012, Bates et al., 2014). Negative regulators of the ACCase in Arabidopsis, BADC1, 2, and 3, have been proposed to function in a competitive manner to inhibit the activity of the heteromeric ACCase (Salie et al., 2016). Another manner of controlling lipid biosynthesis involves the WRINKLED1 (WRI1) transcription factor, which regulates the rate of acyl chain biosynthesis. Several WRI isoforms have been characterized, but their functions seem dependent on their tissue/cell-specific presence. WRI1 stimulates the production of fatty acids, and ectopic 27 expression of WRI1 leads to more oil accumulation in Arabidopsis seeds and seedlings grown on sucrose media (Cernac and Benning, 2004). While WRI1 expression is associated with seeds, WRI3 and 4 activity was narrowed to floral tissues, where WRI4 promoter-GUS fusion-staining also appeared to be present in plant nectaries (To et al., 2012). In seed tissues, WRI1 also controls the expression levels of BCCP2 and thereby ACCase and fatty acid synthesis (Thelen et al., 2001, Ruuska et al., 2002). WRI1 orthologues have been found in different oil storing tissues, for example the embryo in maize (Zea mays), the mesocarp in oil palm (Elaeis guineensis) (Pouvreau et al., 2011, Ma et al., 2013)), castor bean (Ricinus communis), rapeseed (Brassica napus), nasturtium (Tropaeolum majus), the burning bush (Euonymus alatus) (Troncoso-Ponce et al., 2011), and non-seed tissues of Chinese tallow (Triadica sebifera) (Divi et al., 2016). The regulatory nature of WRI1 transcription factors has made them prominent targets for the engineering of oil content in different tissues (Reynolds et al., 2015, Yang et al., 2015). WRI1 alone is not fully capable of turning on TAG biosynthesis, but when co-produced with a diacylglycerol acyltransferase (DGAT), Nicotiana benthamiana leaves were able to achieve higher levels of TAG accumulation than that with WRI1 production alone (Vanhercke et al., 2013). Increases in oil content of Arabidopsis by ectopic production of WRI1 and suppression of starch synthesis resulted in increases of oils in vegetative tissues (Sanjaya et al., 2011). Combining WRI1, DGAT1 and OLE1 ectopic production, while also abolishing sucrose export from leaves and starch synthesis, pushed the flux of carbon to TAG storage even more (Zhai et al., 2017b). In sugarcane, over-expression of WRI1 resulted in doubling of TAG accumulation, but in this plant TAG biosynthesis did not seem to compete with starch biosynthesis unlike in Arabidopsis (Zale et al., 2016). 28 WRI1 is post-translationally regulated through phosphorylation by KIN10 kinase, which is inhibited by increasing levels of sugar, and the phosphorylated WRI1 is targeted for proteolytic degradation (Zhai et al., 2017a, Zhai et al., 2018). Other aspects of WRI1 protein stability have been described, such as a C-terminal PEST motif that leads to destabilization of the protein, and upon removal of the motif to an increase in oil accumulation (Ma et al., 2015). Because storage TAG biosynthesis involves multiple steps across multiple organelles, the full complexity of this process is not yet fully understood, and much can still be learned to realize the full potential of oil production in plant seeds. The balance of eukaryotic and prokaryotic lipid species in Arabidopsis seems to be carefully regulated leading to a finely tuned cellular lipid homeostasis. However, many questions remain about the mechanism by which plants maintain their lipid and acyl composition. For example, it seems likely that there are mechanisms that sense membrane fluidity, acyl saturation levels, and membrane curvature. What are these mechanisms? Some enzymes appear to be sensitive to their membrane environment, providing a potential layer of regulation. For example, though DAG is the preferred substrate for MGD1, the presence of PA and PG appears to be required for MGD1 membrane association and activity (Dubots et al., 2010, Sarkis et al., 2014). Aside from regulation of bulk lipid biosynthesis, some work has been done showing how the degree of plant lipid saturation can be controlled by temperature. The Arabidopsis w-3 fatty acid desaturase FAD8, one of two isozymes present, was shown to be post- translationally regulated through destabilization of the C-terminus when the fad7fad8 double mutant producing a Myc-tagged FAD8 fusion protein was subjected to increasing 29 temperature (Matsuda et al., 2005). More recently, the temperature-dependent stability of ER-localized desaturase FAD3 was shown to be linked to the ubiquitin-proteosome- system, where B. napus and V. fordii FAD3 genes were expressed in yeast and their respective protein levels were modulated by a PEST-like domain at the N-terminus (O'Quin et al., 2010). Lipid Signaling and Response to Stress Lipids are not only integral to the membrane architecture, but they also play an important role in signaling during growth and in times of stress. There is a growing body of work on PA and its role in signaling in response to various cellular conditions. PA and phospholipase D (PLD), which releases PA from other phospholipids, have been implicated in a variety of stress responses by modulating enzyme activity and protein location (Wang, 2005, Ruelland et al., 2015). As in mammalian systems where lipids are found in the bloodstream bound to proteins, PA and other lipid-binding proteins have been identified in the phloem exudates of Arabidopsis (Guelette et al., 2012, Barbaglia and Hoffmann-Benning, 2016) suggesting long-distance movement of lipids through the plant vasculature. In mammalian cells, the importance of PA is apparent through its regulation of small GTPase-activating proteins thereby modulating the activation of Ras signaling pathways (Zhang and Du, 2009). MYB transcription factors interact with PA and their location is influenced by lipid binding. For example, phospholipase D has been implicated in ABA-coordinated stomatal closure, where the PLD product, PA, binds and modulates activity of ABI1 and Sphingosine Kinases (Zhang et al., 2009, Guo et al., 2011, Guo et al., 2012). Furthermore, much like negative feedback regulation of other pathways, 30 PLDa1 producing PA as a product of its lipid hydrolysis has been shown to be itself inhibited by a PA-bound regulator of G-protein signaling, RGS1, which is a GTPase accelerating protein (Roy Choudhury and Pandey, 2017). The role of G-protein signaling is well known in mammalian systems, and plant G-protein signaling is becoming better understood (Pandey, 2017). As more PA-regulated signaling pathways emerge, it is important to begin investigating the mechanisms by which key regulatory enzymes come directly into contact with PA and are regulated by it. Which pools of PA contribute to signaling? Is there negative feedback regulation of the lipid biosynthetic machinery by PA? Is PA or are other lipids critical for enzymatic function of lipid biosynthetic enzymes? Another emerging area of lipid-based regulation of cellular processes focuses on the tight interplay between lipid homeostasis in the plastid and oxylipin biosynthesis. Jasmonic acid (JA), a potent oxidized lipid signaling molecule, is derived from a-linolenic acid in the chloroplast and is active in numerous developmental stages of plant growth as well as involved in responses to stress and herbivory (Howe et al., 2018). Since JA is lipid-derived, various hypotheses exist regarding the identity of the acyl-donating lipid as well as the lipases, which release the α-linolenic acid. However, while the exact molecular mechanism has not yet been elucidated, several lipid mutant phenotypes suggest that particular perturbations of plastid lipid metabolism lead to JA biosynthesis. An interesting example is the dgd1 mutant, which is producing only small amounts of digalactosyldiacylglycerol and as a result, the predominant galactolipid species present in the mutant plant is monogalactosyldiacylglycerol and the mutant is severely stunted in growth (Dörmann et al., 1999). Previously, it was speculated that DGDG was so important that its reduction in content alone would slow down the growth of the plant. However, 31 when crossed with the JA-insensitive mutant coi1, the growth phenotype was rescued (Lin et al., 2016) showing that a disruption in galactolipid biosynthesis also disrupts JA homeostasis leading to the observed growth phenotype. Aside from changes in overall lipid content, JA accumulation has been shown to be triggered by thylakoid-localized lipases (Wang et al., 2018), Plastid LIPases 2 and 3 (PLIP2, PLIP3). The expression of the respective genes is responsive to abscisic acid and the lipases have specificity for PG and MGDG. Thus, the roles that chloroplast membrane lipids and their specific lipases in JA signaling are playing are beginning to come into focus. A more global plant response to stress is the remodeling of membrane lipid composition. An example is the production of oligogalactolipids during freezing in Arabidopsis. During freezing conditions, the activity of SENSITIVE TO FREEZING2 (SFR2) is induced, resulting in the accumulation of oligogalactolipids serving a protective role (Moellering et al., 2010, Moellering and Benning, 2011), since the sfr2 mutants are more sensitive to freezing (Warren et al., 1997). The activity of SFR2 is induced by divalent cations possibly leaking from the chloroplast as a result of membrane disruption in combination with cytosolic acidification during freezing stress (Barnes et al., 2016). Spatial Arrangement and Complexes of Lipid Biosynthetic Enzymes Because changing environmental conditions demand an adjustment of the extent and composition of membranes and the production of lipid-derived signaling compounds, the lipid metabolic machinery must rapidly adjust as well. Much is known about flux through the major lipid biosynthetic pathways, as well as the participating enzymes. However, a knowledge gap remains in understanding the spatial arrangement of lipid enzymes and 32 their protein interactome. The best described lipid complexes are the ACCase, the fatty acid synthase and the TGD complex. While transport of lipids can refer to their inter- compartment movement, the movement of lipids between enzymes located on opposite sides of the same membrane, as well as the translocation of polar lipids between the two leaflets of adjacent membranes needs to be considered as well. For example, plastid PA is formed on the stromal side of the inner envelope membrane (Xu et al., 2006) and MGDG assembly occurs on the leaflet facing the intermembrane space as MGDG synthase is peripherally associated with the intermembrane leaflet of the inner envelope membrane (Xu et al., 2005, Rocha et al., 2016). Given this topology, one has to predict that there is a mechanism that can move PA from the stromal side of the inner envelope membrane to the leaflet facing the intermembrane space where the PA phosphatase is presumably located. It should be noted, that the PA phosphatase has not been definitively localized to a certain leaflet, but its activity is found in the inner membrane and free magnesium ions in the stroma would inhibit a stromal facing phosphatase (Malherbe et al., 1992), suggesting its location at the intermembrane leaflet of the inner envelope membrane. While most of the genes encoding lipid biosynthetic enzymes are known, there is little knowledge on the assembly of the respective proteins into higher order complexes. Plant desaturases appear to function as dimers, with a crystallized diiron D9-destaturase of castor bean providing an example (Schneider et al., 1992, Lindqvist et al., 1996). Recently, the first plant acyl-CoA binding protein structures were solved using rice homologues OsACBP1 and OsACBP2 (Guo et al., 2017). The structure of MGD1 synthase has been described, as well as its PA binding properties (Rocha et al., 2016), 33 though we do not know of any higher oligomeric states. It has been hypothesized that enzymes such as DGD1 and the TGD complex, which also bind PA, could be aggregating as a result of PA binding (Kelly et al., 2016). Though just a hypothesis, these insights pose interesting questions about the mechanisms of regulated oligomerization and possible substrate channeling by lipid enzymes. Moreover, while more data are being collected on enzymatic activities and physiological roles of lipid biosynthetic proteins, the majority of lipid enzyme structures remain to be solved. Overall, more biophysical information is needed on lipid biosynthetic enzymes in order to create accurate models of plant lipid metabolism, identify mechanisms of regulation, and ultimately optimize biofuel and food crops for oil accumulation. Numerous efforts have been made to catalogue and characterize the proteome of plant cells, including chloroplasts (Ferro et al., 2003, Froehlich et al., 2003, Breuers et al., 2011, Lundquist et al., 2017). With this basic information available, the plant lipid community is slowly beginning to assemble the 3D landscapes of our current 2D models depicting lipid metabolism. 34 APPENDIX 35 LPAT G P A T A 1 X A F 4 5 4 5 PA B LPAT GPAT OEM IEM 2 T G D 2 T G D 1 3 1 3 FAS letters are outstanding Figure 2.1. Scheme of phosphatidic acid (PA) synthesis in the plastid and the endoplasmic reticulum (ER). Denoted with transport mechanisms proposed and yet to be characterized: the unknown exporter of acyl groups across the outer envelope membrane (OEM) of the plastid (A) and the transport mechanism by which plastid-assembled PA is moved through the inner envelope membrane (IEM) of the plastid (B). The abbreviations are as follows: fatty acid synthesis (FAS), fatty acid exporter 1 (FAX1), glycerol-3-P acyltransferase (GPAT), lyso- phosphatidate acyltransferase (LPAT), trigalactosyldiacylglycerol 1, 2, 3, 4, 5 (TGD 1, 2, 3, 4, 5). 36 DGD1 DGDG D A G MGD1 MGDG A P A P A P THY MGDG DGDG OEM B IEM Figure 2.2. Proposed movement of galactolipids after synthesis from diacylglycerol (DAG). 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Plant Cell, 21, 2357-2377. 47 CHAPTER 3 A Predicted Plastid Rhomboid Protease Involved in Phosphatidic Acid Metabolism in Arabidopsis thaliana further edited and is published This work has been (https://onlinelibrary.wiley.com/journal/1365313x). Lavell, A., Froehlich, J.E., Baylis, O., Rotondo, A.D. and Benning, C. (2019) A predicted plastid rhomboid protease affects phosphatidic acid metabolism in Arabidopsis thaliana. Plant J, 99, 978-987. in The Plant Journal 48 Abstract The thylakoid membranes of the chloroplast harbor the photosynthetic machinery that converts light into chemical energy. Chloroplast membranes are unique in their lipid makeup, which is dominated by the galactolipids mono- and digalactosyldiacylglycerol (MGDG and DGDG). The most abundant galactolipid, MGDG, is assembled through both plastid and ER pathways in Arabidopsis, resulting in distinguishable molecular lipid species. Phosphatidic acid (PA) is the first glycerolipid formed by the plastid galactolipid biosynthetic pathway. It is converted to substrate diacylglycerol (DAG) for MGDG Synthase (MGD1) which ads to it a galactose from UDP-Gal. The enzymatic reactions yielding these galactolipids have been well established. However, auxiliary or regulatory factors are largely unknown. We identified a predicted rhomboid-like protease 10 (RBL10), located in plastids of Arabidopsis thaliana, that affects galactolipid biosynthesis likely through intramembrane proteolysis. Plants with T-DNA disruptions in RBL10 have greatly decreased 16:3 (acyl carbons : double bonds) and increased 18:3 acyl chain abundance in MGDG of leaves. Additionally, rbl10-1 mutants show reduced [14C]–acetate incorporation into MGDG during pulse-chase labeling, indicating a reduced flux through the plastid galactolipid biosynthesis pathway. While plastid MGDG biosynthesis is blocked in rbl10-1 mutants, they are capable of synthesizing PA, as well as producing normal amounts of MGDG by compensating with ER-derived lipid precursors. These findings link this predicted protease to the utilization of PA for plastid galactolipid biosynthesis potentially revealing a regulatory mechanism in chloroplasts. 49 Introduction Lipid biosynthesis in seed plants is initiated in the chloroplast where de novo fatty acid biosynthesis occurs. Glycerolipids are then assembled at the plastid envelope membranes and the endoplasmic reticulum (ER) (Li-Beisson et al., 2013). Due to the distinct specificity of acyltransferases associated with the different membranes, the lipid species from each compartment have either a 16-carbon acyl chain at the sn-2 position (plastid) or an 18-carbon acyl chain (ER) (Browse et al., 1986, Mongrand et al., 1998). Phosphatidic acid (PA) is the initial glycerolipid to be assembled and then converted to other lipids. Through the use of both the chloroplast and ER lipid precursors for galactolipid biosynthesis, Arabidopsis thaliana maintains a specific 16:3/18:3 ratio of acyl chains (acyl carbons : double bonds) in its chloroplast MGDG molecules. We know the genes that encode the PA biosynthetic enzymes in the chloroplast that first acylate glycerol-P and then lysophosphatidic acid (Kunst et al., 1988, Nishida et al., 1993, Xu et al., 2006) as well as the genes encoding the galactosyltransferase MGD1, which uses diacylglycerol (DAG) (Jarvis et al., 2000, Awai et al., 2001, Rocha et al., 2016) derived from dephosphorylation of PA by a phosphatase. However, the gene encoding the PA phosphatase responsible for creating the bulk plastid DAG pool in vegetative tissues has only been tentatively identified (Nakamura et al., 2007) and its topology within the inner envelope membrane has yet to be determined, although its activity has been measured in the chloroplast envelopes (Block et al., 1983, Malherbe et al., 1992). The likely location of the plastid PA biosynthetic enzymes on the inside of the inner chloroplast envelope membrane and the presence of the galactolipid biosynthetic enzyme MGD1 on the outer leaflet of the inner chloroplast envelope membrane (Xu et al., 2005) poses the question 50 for the movement of PA and its conversion product as well as MGD1 substrate, DAG, through the inner envelope membrane. Due to its polarity, PA cannot freely flip between the membrane leaflets and a PA transporter may have to be invoked. In any case, the precise mechanism for providing MGD1 with prokaryotic substrate DAG for the biosynthesis of MGDG is not yet known. On the contrary, a likely mechanism for ER- derived PA insertion into the inner envelope membrane is through the action of the TGD complex, which is a multiprotein ABC transporter, TGD1, 2, 3, proposed to transport ER- assembled PA to the inner envelope membrane of the chloroplast (Xu et al., 2005, Roston et al., 2012, Yang et al., 2017). TGD2, one component of the TGD1, 2, 3 complex contains a PA binding site and is physically located in the inner envelope membrane spanning the intermembrane space between the two envelope membranes (Awai et al., 2006, Lu and Benning, 2009, Roston et al., 2011). TGD4 is a PA-binding protein in the outer envelope membrane also involved in the process (Wang et al., 2013). However, it should be noted that it has not yet been directly shown that PA is actually transported by the TGD proteins. Thus, while we have a reasonable understanding of how lipid precursors produced at the ER are translocated to the inner envelope membrane and utilized for galactolipid biosynthesis, it is still unclear how chloroplast PA biosynthesis contributes plastid-derived precursors for galactolipid biosynthesis. Identifying genes encoding lipid transporters that could be involved in PA transfer from the stroma side to the intermembrane face of the inner envelope membrane may provide additional targets for the regulation of lipid assembly. One mechanism of regulation yet to be implicated in plant lipid metabolism is intramembrane proteolysis. We identified a candidate gene (At1g25290) encoding a 51 predicted intramembrane protease, rhomboid-like protein (RBL10) that appears to be involved in the regulation of chloroplast PA metabolism and transport. Rhomboid-like protein 10 (RBL10) belongs to a superfamily of intramembrane proteases called rhomboids, with the founding member discovered in Drosophila melanogaster (Bier et al., 1990). Members of this superfamily are multi-pass integral serine proteases found in nearly all organisms, and they typically serve regulatory roles. Aside from directly initiating a signaling cascade, as in the case of Rhomboid-1 and the epidermal-growth factor pathway (Lee et al., 2001, Urban et al., 2001), rhomboids can also influence assembly of complexes, for example Providencia stuartii rhomboid AarA, which processes TatA and directs the oligomerization of the twin arginine translocase complex used in quorum sensing (Stevenson et al., 2007). In humans, rhomboids have been implicated in Parkinson’s disease by maintaining mitochondrial integrity through proteolysis of Pink1 by the mitochondrial PARL protein, which affects Pink1 trafficking (Meissner et al., 2011, Meissner et al., 2015). Much like in mice and animals, 13 rhomboids have been predicted to exist in Arabidopsis (Lemberg and Freeman, 2007, Page and Di Cera, 2008), and although they have been compared to characterized animal rhomboids (Kanaoka et al., 2005), a detailed molecular mechanism for any of them has yet to be described. Of the two rhomboid-like proteins predicted to be located in the chloroplast of Arabidopsis, only RBL10 seems to be involved in lipid metabolism based on our own observations. The second plastid rhomboid protease, RBL11, may have other functions that do not relate to lipid homeostasis. Here we describe in detail the analysis of rbl10 mutants and propose how an integral membrane protease may be involved in the metabolism or transport of PA in the Arabidopsis chloroplast. 52 Results and Discussion Discovery of a Role for RBL10 in Chloroplast Lipid Metabolism To investigate possible factors required for metabolism and trafficking of PA assembled in the chloroplast, the Chloroplast Phenomics 2010 database (Ajjawi et al., 2010) was searched for T-DNA mutants showing a shift in the 16/18 carbon ratio in total leaf acyl composition. We predicted that reduced utilization of plastid PA would result in an overall decrease in 16:3 acyl chain abundance. One putative mutant was identified, and the gene disrupted in the T-DNA mutant line encodes a rhomboid-like protein located in the chloroplast (Fig. 3.1a). To corroborate that a mutation in this gene resulted in the observed lipid phenotype, we obtained additional T-DNA insertion mutants for RBL10 (Fig. 3.1a). They showed decreased RBL10 transcript abundance when amplifying a cDNA segment by qPCR (Fig. 3.1b). Analyzing the acyl composition of individual lipids using thin-layer chromatography (TLC) in combination with gas-liquid chromatography (GLC) it became apparent that the primary lipid bearing the acyl changes seen in the total lipid FA profile is MGDG (Fig. 3.1c). The ratio of 16:3 to 18:3 acyl chains in rbl10-1, rbl10- 2, and rbl10-3 was reduced several fold, suggesting a predominance of ER precursors contributing to MGDG biosynthesis. Though each line tested is disrupted in a different location of the RBL10 gene, all showed similar MGDG acyl compositional changes. Furthermore, the overall amount of MGDG in the mutant lines was slightly reduced compared to wild type but not statistically significant (Fig. 3.S1a). The changes in MGDG acyl composition were restored to wild-type levels by reintroducing an RBL10 coding sequence into the different mutant alleles under the control of the 35S promoter (Fig. 53 3.1d). Based on this result we concluded that a disruption in RBL10 is the primary cause for the observed change in acyl composition in MGDG. Since the chloroplasts of Arabidopsis appear to have a second predicted rhomboid-like protease, three independent T-DNA insertion lines disrupting RBL11 (At5g25752) were analyzed for their total and MGDG acyl compositions. No changes in the acyl composition of rbl11 mutants were observed (Fig. 3.S2) and the analysis of this mutant was not further pursued in the context of this study. JA and JA-Ile Levels are Affected in rbl10 The changes seen in MGDG composition did not affect the overall plant appearance or growth under the standard lab conditions tested. Because RBL10 had tentatively been implicated in jasmonic acid (JA) signaling or metabolism, we quantified this hormone (Thompson et al., 2012). Upon leaf wounding, the rbl10-1 and rbl10-2 mutant lines accumulated significantly less JA and the isoleucine derivative of JA, JA-Ile, at 20 minutes compared to wild type (Fig. 3.S3a, b) (0.006 ³ p, 0.004 ³ p and 0.044 ³ p, 0.064 ³ p respectively). The amount of the JA precursor 12-Oxophytodienoic acid (OPDA) accumulated by the two lines at 20 minutes past wounding was not as reduced when compared to wild type (Fig. 3.S3c) (0.081 ³ p, 0.090 ³ p). These changes are indicative of an attenuation of JA biosynthesis. Aside from generally normal vegetative growth under standard growth chamber conditions, it was previously reported that rbl10 mutants have altered pollen morphology, floral defects along with a reduced number of seeds filled per silique, and longer roots with more side branching (Thompson et al., 2012). An attenuated biosynthesis of JA and JA-Ile could explain some of these reported phenotypes. In our 54 own work, we were able to corroborate the reduced number of seeds per silique of rbl10- 1 as compared to wild type (Fig. 3.S4). At this time, the pollen, seed, and hormone phenotypes cannot be directly linked to altered galactolipid metabolism. However, Thompson et al. (Thompson et al., 2012) suggested that possible disruptions of JA signaling might be the cause of the observed developmental changes in the rbl10 mutants and there is an increasing body of work demonstrating a relationship between chloroplast glycerolipid metabolism and JA biosynthesis (Lin et al., 2016, Wang et al., 2018). RBL10 is Located in the Inner Envelope Membrane of Chloroplasts The predicted structure of the RBL10 transcripts of Arabidopsis suggests two splice forms as shown in Figure 1a. Previously, the subcellular targeting of RBL10 to the chloroplast was demonstrated with fluorescent tagging and chloroplast fractionation (Knopf et al., 2012, Thompson et al., 2012). Here, we further identified into which envelope membrane RBL10 is inserted. In-vitro translated RBL10.1 and RBL10.2 proteins labeled with 3H- Leucine were imported into pea chloroplasts. Fractionation experiments showed that both proteins are present in the chloroplast envelope membranes (Fig. 3.2a). To more precisely evaluate the subcellular localization of RBL10, we performed import and protease protection assays (Fig. 3.2b). We showed that imported RBL10.1 and 10.2 proteins are resistant to trypsin cleavage (Fig. 3.2b) unlike Toc33, a known outer envelope membrane-resident protein. Taken together, we conclude from these results that both versions of RBL10 specifically insert into the inner envelope membrane of the chloroplast. A homology model produced with SWISS protein using the crystal structure of GlpG, which has a 27% amino acid identity with RBL10, (Wang et al., 2006), suggested 55 how the six transmembrane helices of RBL10 are oriented in the membrane (Fig. 3.2c), with the predicted active site serine and histidine residues close to the leaflet solvent interface. Since rhomboids are membrane-bound proteases, their substrate must be inserted into the same membrane in order for proteolysis to occur (Moin and Urban, 2012, Guo et al., 2016). However, at this time a substrate protein remains to be identified that either directly or indirectly affects lipid metabolism. ER-Assembled Lipids Serve as Primary Precursors for MGDG Biosynthesis in the rbl10-1 Mutant To probe the biochemical basis of the observed changes in MGDG acyl composition, pulse-chase radiolabeling experiments were performed. Detached leaves were floated on [14C]–acetic acid-containing buffer, which resulted in the incorporation of label into lipid acyl groups by de novo fatty acid synthesis. During the first hour after introducing the radiolabel (pulse), chloroplast lipid assembly was observed by lipid profiling using TLC of lipid extracts from the leaves and determining the incorporation of radiolabel into polar lipids (Fig. 3.3). Comparing the wild type and the rbl10-1 mutant, less than half of the radiolabel was incorporated into MGDG in rbl10-1. The order (from high to low) of levels of incorporation of label into polar lipids during the pulse in wild type was MGDG, phosphatidylcholine (PC), phosphatidylglycerol (PG), PA, phosphatidylinositol (PI)/ phosphatidylethanolamine (PE), sulfoquinovosyldiacylglycerol (SQDG), and DGDG respectively. In the rbl10-1 mutant, this order was changed (from high to low): PC, PG, MGDG, PA, PI/PE, SQDG, and DGDG. In addition to the reduction of label in MGDG in rbl10-1, the level of label in PA was equal to that of MGDG at 20 minutes. In comparison, 56 wild-type leaves accumulated label in MGDG to a level three-fold of that accumulated in PA by 20 minutes. The chase, after the leaves were moved to new buffer containing only unlabeled acetate, showed no major differences in rates of turnover between wild-type and rbl10-1 leaves. Because there was label accumulating in PA and PG, as well as PC, we concluded that the chloroplast pathway is functional up to the step just prior to MGDG biosynthesis. PA Synthesized in Chloroplasts is Used for PG Biosynthesis but not MGDG Since MGDG biosynthesis can occur from both plastid and ER lipid precursors, [14C] – acetate feeding of intact, isolated chloroplasts should allow us to focus on only the plastid pathway reactions. In wild-type intact chloroplasts, radiolabel accumulated to the same levels in both PA and MGDG by 5 minutes, with MGDG accumulating the most label at all time points after 5 minutes (Fig. 3.4a). Intact rbl10-1 chloroplasts accumulated three times as much radiolabel in PA than in MGDG by 5 minutes, and MGDG never accumulated more label than PA or PG (Fig. 3.4a). Based on the TLC plate image (Fig. 3.4b), the differences in label accumulation between wild-type and rbl10-1 chloroplast lipids were striking, corroborating the data observed during the whole leaf pulse-chase experiment. Figure 3.5a shows the different fates of PA in the chloroplast. Our interpretation of the labeling results was that MGDG biosynthesis from the chloroplast-derived precursor lipid PA is compromised in rbl10-1 mutant plants (Fig. 3.5aii, iii) while the biosynthesis of PG proceeds normally (Fig. 3.5ai). 57 MGD1 Activity is not Affected by Loss of RBL10 While rbl10-1 chloroplasts were not able to synthesize MGDG from de novo assembled PA, these chloroplasts were not compromised in MGD1 enzymatic activity when supplied with DAG and [14C] – UDP-galactose (Fig. 3.5B). The ability of rbl10-1 chloroplasts to utilize exogenous MGD1 substrates along with only slightly reduced overall MGDG levels, but altered acyl composition in the rbl10 mutant plants, supports the hypothesis that there is a shift in in the mutants in the utilization of lipid precursors to predominantly ER lipid precursors for MGDG biosynthesis. This shift has also been previously observed for the ats1 mutant disrupted in the chloroplast glycerol-3-phoshate acyl transferase leading to the formation of lysophosphatidic acid (Kunst et al., 1988). However, in the ats1 mutant, the effect was less specific to MGDG and chloroplast PG levels are reduced as well. This again speaks for a disruption after the formation of chloroplast PA but prior to MGDG assembly in the rbl10-1 mutant. Therefore, we tested the activity of PA phosphatase activity next. PA Plastid Phosphatase Activity is Increased in rbl10-1 Mutants To further narrow the possible protein targets affected by the disruption of the RBL10 gene (Fig. 3.5a), we tested the activity of the PA phosphatase by using [14C]–PA and mixed inner and outer envelope membranes from wild-type and rbl10-1 chloroplasts. To our surprise, the envelope membranes of rbl10-1 chloroplasts dephosphorylated PA to diacylglycerol at a higher rate than the wild-type membrane preparations, with 11% and 8% conversion at 30 minutes respectively (Fig. 3.5c). Our initial hypothesis was an impaired conversion of PA to MGDG in chloroplasts due to a reduction in PA phosphatase 58 activity. However, we concluded based on our data that the changes in plastid PA utilization was not due to compromised PA phosphatase activity. Plastid PA is synthesized by two acyltransferases, ATS1 present in the stroma (Xu et al., 2006) and ATS2 likely associated with the stroma side of the inner envelope membrane (Yu et al., 2004). Therefore, we propose that the physical translocation of PA from its place of synthesis by the two acyltransferases ATS1 and ATS2 on the inner leaflet of the inner envelope membrane to its intermembrane-facing leaflet, where presumably the PA phosphatase and, for certain, the MGD1 synthase are located, is disrupted in the rbl10-1 mutant. A Refined Model of PA Metabolism in Arabidopsis Chloroplasts To describe this system, it seems instructive to draw parallels with the ER phospholipid transporter ALA10, which is reported to form a complex with the ER desaturases FAD2/FAD3 to modulate the balance between PC and MGDG (Botella et al., 2016). Hence, it seems plausible that a PA transporter could be in complex with the plastid PA biosynthetic enzymes and possibly even the PA phosphatase, resulting in substrate channeling. In our refined model (Fig. 3.6), we accommodate our current knowledge (Hurlock et al., 2014) about the transport of ER–derived PA from the ER through the outer to the inner envelope membrane by the TGD complex (Fig. 3.6a). In addition, we hypothesize that a not yet identified transporter assists in the transfer of chloroplast- assembled PA from the inside of the inner envelope membrane, where it is synthesized, to its intermembrane space-facing leaflet, where it is converted to DAG and ultimately MGDG (Fig. 3.6b). There are currently at least three predictions based on this hypothesis 59 that need to be satisfied for the model to be correct: 1. The plastid PA phosphatase must be associated with the intermembrane space-facing leaflet of the inner envelope membrane, which would give it access to ER-derived and chloroplast-derived PA precursors for DAG and ultimately MDGD biosynthesis. 2. A transporter must exist that either can flip or channel chloroplast assembled PA through the inner envelope membrane. 3. RBL10 must affect the assembly or insertion of this transporter and associated proteins in the inner envelop membrane. Aside from testing the hypothesis outlined above, it will be important to actually demonstrate that RBL10 acts as an intramembrane protease. Reconstituting and assaying a six-transmembrane integral protein is challenging. Moreover, even though numerous synthetic rhomboid substrates have been identified and can serve as substrates for diverse rhomboids, finding true in vivo substrates for rhomboids has not been trivial in the past. Even for the well-studied rhomboid, GlpG from E. coli, the only crystallized rhomboid (Wang et al., 2006), the native substrate remains unknown. Absent of our knowledge of a native RBL10 substrate, it will be difficult to distinguish whether RBL10 directly processes components of a PA-transport/synthesis complex in the inner chloroplast envelope membrane, or whether it releases a signaling peptide that could indirectly affect this process. The list of potential protein interactors/substrates of RBL10 is vast, even though a previous proteomic analysis of an RBL10/RBL11 double mutant (Knopf et al., 2012) may provide a starting point. An intriguing feature of rhomboids is the nature of their substrate recognition, which is determined by the topology of the substrate in the membrane rather than a specific protein sequence (Urban and Freeman, 2003). Even though rhomboids do not recognize 60 a consensus sequence, their substrate recognition mechanism offers surprising specificity for their transmembrane helix substrates. Moreover, rhomboids scan for their substrate moving through membranes at biologically relevant rates due to the nature of their interaction with the membrane (Kreutzberger et al., 2019). Unfortunately, the lack of a concrete cleavage consensus site poses a challenge when searching for an endogenous substrate and ultimately a molecular mechanism to explain the observed rbl10 lipid phenotypes. However, our current data point towards a testable working hypothesis that should allow us to narrow down the substrate for RBL10 in the future. Materials and Methods T-DNA Lines Seeds for T-DNA lines used were obtained from the Arabidopsis Biological Resource Center (ABRC). The lines SALK_036100, SALK_037037, and SALK_092293 were denoted as rbl10-1, rbl10-2, and rbl10-3 in this study respectively. Plant Growth Conditions Seeds were sown directly into soil for lipid analysis and pulse-chase labeling. Plants were grown in a growth chamber at 22°C, a 16-hour light/8-hour dark cycle, and with approximately 120 μmol of light. For isolation of chloroplasts and envelopes, Arabidopsis seeds were sterilized and grown on Murashige and Skoog (MS) agar-solidified medium in Percival growth chambers at 22°C, a 16 hours light/8 hours dark cycle, and 120 μmol of light. 61 DNA Constructs and Complementation The RBL10, splice variant AT1G25290.2, coding sequence was amplified from cDNA. Plant RNeasy Mini Kit (Qiagen, www.qiagen.com) was used to extract mRNA of Col-0 plants including the optional on-column DNase digestion, following reverse transcription with SuperScript III Reverse transcriptase (Invitrogen, www.thermofisher.com) and its corresponding first strand synthesis protocol, excluding RNaseOUT (recombinant rnase inhibitor). Full length coding sequence excluding stop codon was amplified using Phusion High-Fidelity DNA polymerase (Thermo Scientific, www.thermofisher.com) and gel- purified. The amplicon was directionally inserted into a Gateway pENTR/D/TOPO entry vector (Invitrogen) and sequence verified vector was linearized using PvuI restriction enzyme (New England Biolabs, www.neb.com) and gel purified. The linear vector was recombined with the pEARLEYGATES101 expression vector using the gateway LR Clonase II kit (Invitrogen). The resulting construct was introduced into Agrobacterium tumefaciens GV3101 and used to complement the rbl10-1 mutant plants using the floral dip method and BASTA (glufosinate-ammonium, BASF, www.basf.com) selection (Logemann et al., 2006). Homology Modeling Swiss Model software was used to generate a homology model of RBL10 (Biasini et al., 2014) based on the published crystal structure of GlpG (E. coli) (Dickey et al., 2013) deposited at the Protein Data Bank ( 4NJN). Protein structure further processed in PylMol. 62 Intact Chloroplast Isolation Arabidopsis seedlings were plate-grown on agar-solidified MS medium in a dense lawn and harvested at 3-weeks of age and intact chloroplasts were isolated following the protocol in (Cline and Keegstra, 1983) with modifications. Plant tissue was homogenized with a polytron in a grinding buffer at pH7.3 (330mM Sorbitol, 50mM HEPES, 1mM MgCl2, 1mM MnCl2, 2mM EDTA, 0.1% BSA), filtered through miracloth, and crude chloroplasts were pelleted at 700xg. The pellet was resuspended and loaded on a continuous Percoll gradient (generated by centrifugation at 39,000 x g, 30 mins, no brake). Intact chloroplasts were collected from the lower phase of the gradient after centrifugation at 2620 x g for 10 mins without braking. Intact chloroplasts were washed twice in an import buffer of pH 8.0 (330mM Sorbitol, 50mM HEPES) and yield approximated by chlorophyll equivalents. Mixed envelopes were isolated by rupturing purified intact chloroplasts as above using a Dounce homogenizer (Wheaton 15mL) in TE (10mM Tricine pH 7.5, 2mM EDTA) buffer with 0.6M sucrose. Thylakoid membranes were separated by centrifugation of the lysed chloroplast solution at 1,500xg for 5 minutes, repeated 3 times by transferring the supernatant to a new tube each time. The entire volume of supernatant was combined and centrifuged at 100,000xg for 1 hours to collect the remaining inner and outer membranes. The resulting pellet of envelope membranes was then resuspended in approximately 100 μL of PAP assay buffer and the protein content of each prep was quantified using an RCDC protein quantification kit (Bio-rad). For in vitro import assays, intact pea chloroplasts were isolated from 8 to 12 day- old pea seedlings (Little Marvel, Dwarf Variety, Livingston Seed Co., Columbus, OH 43216) and purified over a Percoll gradient as previously described (Bruce, 1994). Intact 63 pea chloroplasts were re-isolated and resuspended in import buffer (330 mM sorbitol, 50 mM Hepes/KOH, pH 8.0) at a concentration of 1 mg chlorophyll per mL. In Vitro Translation of precursor proteins Precursor proteins used in import assays were radiolabeled using [3H]-Leucine (approximately 0.05mCi/50μl Translation Reaction is used) and translated using Promega’s TNT® Coupled Reticulocyte Lysate System according to the manufacturer’s protocol. After translation, labeled precursors were diluted with an equal volume of ‘cold’ 50mM L-Leucine in 2X Import Buffer (IB). Import and Protease Protection Assays Import assays were performed essentially as described in Tripp et al. (Tripp et al., 2007). Essentially: 100 μl chloroplasts (1mg chlorophyll/ml), 4 mM Mg-ATP/IB final concentration, and 100 μl radio-labeled precursor protein to a final volume of 300 μl was incubated for 30 minutes at room temperature, under room light. After import, the reaction was divided into two 150 μl aliquots. One portion was not further treated with protease [(- ) control] and intact chloroplasts were directly recovered by centrifugation through a 40% Percoll cushion. The other portion was incubated with Trypsin as previously described (Tripp et al., 2007) for 30 minutes on ice. After quenching Trypsin with Trypsin Inhibitor, chloroplasts were again recovered by centrifugation through a 40% Percoll cushion, lysed and then fractionated into a total soluble (S) and total membrane fraction (P). 64 Fractionation of Imported RBL10 The membrane association of RBL10.1 and RBL10.2 was determined by performing a large-scale import assays and subsequent fractionation experiments. Briefly, either [3H]- Leucine labeled RBL10.1 or RBL10.2 were incubated with chloroplasts using a large- scale version of our import assay. After import, intact chloroplast were recovered, lysed and applied to a sucrose step-gradient according to the method of Li et al. (Li et al., 2017) and fractionated into envelope membranes (E), stroma (S) and thylakoid membranes (T). All fractions were analyzed by SDS-PAGE and fluorography. Controls: Toc33, outer envelope membrane protein; FtsH8, thylakoid membrane protein and mature small subunit of Rubisco (mSSU), stroma protein. Note: The RBL10.1 and RBL10.2 fluorograms were exposed to film 3X longer than control samples. Lipid Analysis Lipids were extracted from detached Arabidopsis rosette leaves as well as isolated chloroplasts and prepared for Gas-Liquid Chromatography (GLC) following Wang et al, 2011 (Wang and Benning, 2011). Polar lipids were separated using a silica TLC plate treated with (NH4)2SO4 and an acetone:toluene:water (91:30:7 v/v) solvent system. Lipid bands were visualized with brief iodine vapor staining. Individual lipids were scraped, and their fatty acid profiles analyzed using GLC. Composition is presented as a mole percentage of total fatty methyl esters detected in each lipid. 65 Radiolabeling Pulse-chase labeling was performed with detached four-week old Arabidopsis leaves floated on 25mM 4-Morpholineethanesulfonic acid hemisodium salt (MES-KOH) (pH 5.7), 0.01% TritonX-100 buffer. A table top light (25 Watts at 5.5 cm distance) was used to stimulate lipid biosynthesis. De novo fatty acid synthesis was observed by pulsing with 120 μCi of [1,2-14C] - acetic acid sodium salt (ARC 0173 - American Radiolabeled Chemical, Inc.) for an hour, washing the leaves with same MES buffer without radiolabel twice, and the chase was performed by continuing to incubate leaves in the MES buffer without radiolabel. At each time point measured, a single leaf was taken, and lipids were extracted for TLC separation. Incorporation of radioactivity was imaged/measured by exposing a Kodak phosphoscreen with the TLC plate and scanning the screen with Quantity One software. Quantification of radioactivity in lipids was also performed by scraping the silica off the TLC plate containing the lipid band and counting using a Liquid Scintillation Counter (LSC) and 4a20 counting cocktail (Research Products International, www.rpicorp.com). Isolated Arabidopsis chloroplasts were fed with [1,2-14C] - acetic acid (ARC 0173, American Radiolabeled Chemical, Inc.) to measure plastid lipid biosynthesis only. Chloroplasts were exposed to table top light to stimulate lipid synthesis, aliquots were taken from each sample at various time points and placed directly into lipid extraction buffer. Lipids were separated with TLC and quantified with a LSC and Quantity One Software. Intact Arabidopsis chloroplasts were isolated and fed [1-14C] – UDP- galactose (ARC 3472, American Radiolabeled Chemical, Inc.) to measure plastid MGD1 activity. 66 About 0.30 μCi of activity per 220 μg of chlorophyll equivalents was used. Chloroplasts were resuspended in assay buffer pH7.6 (330mM Sorbitol, 50mM Hepes, 1mM MgCl2, 1mM MnCl2, 5mM EDTA) with 100 μM DAG (1,2-Dioleoyl, Sigma Aldrich D0138), exposed to table top light to stimulate lipid synthesis, and aliquots were taken from each sample at 3,10 and 30 minute time points and placed directly into lipid extraction buffer. Lipids were extracted and separated with polar TLC as described in lipid analysis section above, and then quantified with LSC and Quantity One. Arabidopsis mixed chloroplast envelopes were used to assay for phosphatidic acid phosphatase activity. Equal amounts of protein were incubated with di[oleoyl-1-14C] - phosphatidic acid (ARC 1303, American Radiolabeled Chemical, Inc.). Envelope preparation done as described in this study and the PAP assay was performed as in (Xu et al., 2005). Lipids were extracted and separated with TLC and radioactivity in TAG was quantified by LSC. Acknowledgement Anna Hurlock, Dr. Patrick Horn, Kun Wang, and Dr. Yang Yang provided advice and useful discussions that aided this work. Dr. John Froehlich performed the chloroplast import assays, contributing to Figure 3.1. Financial support for this work was primarily from the Division of Chemical Sciences, Geosciences and Biosciences, Office of Basic Energy Sciences of the United States Department of Energy (Grant DE-FG02- 91ER20021). In addition, Anastasiya Lavell was supported in part by a predoctoral training grant from the National Institute of General Medical Sciences of the National Institutes of Health (Grant Number T32-GM110523). 67 APPENDICES 68 APPENDIX 3A Main Text Figures 69 (a) AT1G25290.1 AT1G25290.2 1 rbl10-2 rbl10-3 rbl10-1 9 1985bp 2 3 4 5 6 7 8 (b) WT rbl10-1 rbl10-2 rbl10-3 l e g n a h C d o F A N R m 1.0 0.8 0.6 0.4 0.2 0 N.D. N.D. WT rbl10-1 rbl10-2 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 WT rbl10-1 rbl10-1:RBL10 (1) rbl10-1:RBL10 (2) rbl10-1:RBL10 (3) rbl10-1:RBL10 (4) (c) G D G M f o % l o M (d) G D G M f o % l o M 80 60 40 20 0 80 60 40 20 0 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 Acyl Groups Figure 3.1. Structure and phenotypes of rbl10 mutant alleles. (a) Gene model of RBL10 and the two splice forms (At1g25290.1 and At1g25290.2), where in At1g25290.2 exon 6 has 21 fewer nucleotides at the exon’s 5’ end. Positions of the T-DNA insertions in the mutant lines used in this study are shown on the gene model. (b) Quantitative PCR showing reduction in RBL10 gene transcript abundance amplified from cDNA prepared from the rbl10-1 and rbl10-2 knock-out mutants (closed bar) or wild type (WT) (open bar) n=4. (c) Acyl composition of MGDG (carbon # : double bond #) measured in the three rbl10 alleles and WT, showing decreased 16:3 and increased 18:3 relative acyl chain abundance in the mutants n=3. (d) WT acyl chain composition of MGDG restored by expression of the RBL10 coding sequence driven by a 35S promoter in the rbl10-1 background n=3. Error bars report SE but are in most cases too small to show. 70 (a) RBL10.1 RBL10.2 FtsH8 Toc33 mSSU (c) Fractionation E S T (b) MW 37- RBL10.1 + TP P S P S - TR p MW 100- Controls - + TP P S P S TR p p m m 25- MW 37- RBL10.2 + TP P S P S - TR p 1 2 3 25- m 1 2 3 4 m Arc6 75- 37- FtsH8 p m Toc33 1 2 3 4 Intermembrane Space His-293 Ser-240 Chloroplast Stroma Figure 3.2. Localization of RBL10 to the Inner Envelope Membrane of Chloroplast. (a) The RBL10 protein was labeled with [3H]-Leucine, imported into chloroplast and subsequently fractionated into various subcellular compartments. The presence of the protein exclusively in the envelope “E”, but not in the stroma “S” or thylakoid “T” fractions confirms its localization. (b) Import and protease protection assays further confirmed that RBL10 was specifically localized to inner envelope membrane. Resistance of imported RBL10 to trypsin (+) digestion is indicated by the presence of a band in the pellet fraction “P”, thus confirming that RBL10 is deeply imbedded into the inner envelope membrane. (c) Homology model of RBL10 made with SwissModel using the GlpG crystal structure as a template (PDB 4njn.1.A). Membrane boundaries are shown for illustration purposes only, predicted catalytic serine and histidine residues are shown in yellow and green respectively. 71 PA 45 PG DGDG SQDG PI/PE PC MGDG WT s d p i i l r a o p l n i l l e b a o d a r i % s d p i i l r a o p l n i l l e b a o d a r i % 30 15 0 15 0 0 20 45 30 15 0 15 40 (min) 60 0 5 10 15 (hr) 20 25 rbl10-1 0 0 20 40 (min) 60 0 5 10 (hr) 15 20 25 Figure 3.3. Radiolabeled pulse chase of detached leaves using 14C-acetic acid. A decrease in incorporation of radioactivity into MGDG and increased incorporation into PG and PA of the rbl10-1 mutant (bottom panels) compared to the wild type (WT, top panels) is shown n=3. Error bars report SE but are often smaller than the symbol. Abbreviations: DGDG, digalactosyldiacylglycerol; MGDG, monogalactosyldiacylglycerol; PA, phosphatidic acid; PC, phosphatidylcholine; PG., phosphatidylglycerol; PI/PE, phosphatidylinositol/phosphatidylethanolamine; SQDG, sulfoquinovosyldiacylglycerol. 72 (a) s d p i i l r a o p l n i l l e b a o d a r i % 50 40 30 20 10 0 (b) PA MGDG PG PC WT 70 60 50 40 30 20 10 rbl10-1 0 5 10 15 (min) 0 0 20 5 10 15 (min) 20 WT rbl10-1 PA MGDG PG DGDG 5 10 15 20 5 (min) 10 15 20 Figure 3.4. Labeling of isolated chloroplasts of rbl10-1 mutant and WT with 14C- acetate. (a) Labeling time course with isolated chloroplasts of rbl10-1 (right panel) showing an increased 14C-acetate accumulation into PA with an apparent conversion of PA into PG compared to wild type (WT, a, left) which appears to convert PA to MGDG n=3. Error represent SE and are often smaller than the labels. (b) TLC image of the extracted lipids of the chloroplasts labeled with 14C-acetate in (a), one representative of three. The accumulation of radiolabel into MGDG of WT chloroplasts is approximately twice the accumulation of radiolabel into MGDG for rbl10-1 chloroplasts. The chloroplasts of both mutant and WT appear to be capable of converting PA to PG. Abbreviations of lipids as in the Figure 3 legend. 73 PA ii i DAG MGDG iv DGDG rbl10-1 WT MGDG_WT MGDG_rbl10-1 DGDG_WT DGDG_rbl10-1 iii (c) G A D n i l 12 10 8 6 0 l e b a o d a r i % (a) (b) l l PG 600 500 400 300 200 100 0 0 y h p o r o h c l / g u M P C 10 (min) 20 30 10 20 (min) 30 Figure 3.5. Probing PA metabolism in rbl10-1. (a) Overview of plastid PA metabolism. PA can be converted to PG (ai) or is dephosphorylated to supply diacylglycerol (DAG) pool (aii) for galactolipid biosynthesis (aii, iv). It is important to note that DGDG biosynthesis favors precursors from the ER pathway not shown here (aiv). (b) Activity of the galactolipid enzymes MGD1 and DGD1 in isolated chloroplast using labeled UDP-Gal as precursor. They synthesize MGDG and DGDG respectively and are both functional at wild-type (WT) levels in rbl10-1 isolated chloroplasts n=3. (c) Phosphatidic acid phosphatase activity of envelope membranes supplied with 14C-PA n=3. While rbl10-1 chloroplasts cannot readily convert de novo synthesized PA to MGDG, mixed envelopes of this mutant showed an elevated PA phosphatase activity. Error bars report SE. Error bars are often smaller than the symbols. 74 TGD 2 TGD 2 1 3 1 3 PA PAP MGD1 DAG (a) 4 5 4 5 (b) 4 5 4 5 TGD 2 TGD 2 1 3 1 3 ? PA OEM MGDG IEM OEM MGDG IEM PAP DAG MGD1 LPAT GPAT FAS Figure 3.6. Proposed model of PA metabolism in the chloroplast envelope membranes. (a) Fate of PA assembled at the ER and imported back to the plastid. (b) Fate of PA assembled and metabolized in the plastid. Both panels depict ER and plastid PA serving as a substrate for the same PA phosphatase in the intermembrane space leaflet of the inner membrane of the chloroplast. The import of ER sourced PA is hypothesized to be facilitated by the TGD complex in the plastid outer and inner envelope membranes (a). The plastid PA is assembled on the stromal side of the inner envelope membrane. A transport mechanism is proposed, which translocates PA and inserts it into the leaflet of the inner membrane facing the intermembrane space (b). The rbl10 mutant phenotype can be explained assuming that RBL10 is critical for the function of the inner envelope PA transporter. 75 APPENDIX 3B Supplementary Figures 76 (b) WT rbl10-1 rbl10-2 % l o M (d) MGDG PG DGDG SQDG PC PG 60 50 40 30 20 10 0 80 60 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 SQDG (a) 40 30 % l o M 20 (c) 10 0 40 30 20 % l o M (e) % l o M 10 0 60 50 40 30 20 10 0 Total 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 DGDG 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 PC 40 % l o M 20 0 40 30 (f) % l o M 20 10 0 77 16:0 16:1 16:2 18:1 18:2 18:3 16:0 16:1 16:2 16:3 18:0 Acyl Groups 16:3 18:0 Acyl Groups 18:1 18:2 18:3 Figure 3.S1. Full lipid analysis of wild-type (WT) and rbl10 leaves. (a) Polar lipid profile of WT, rbl10-1 and rbl10-2 leaves as a mole percentage of the measured polar lipids. Although MGDG appears slightly reduced in the mutant lines, this difference is not significant at the 0.05 level based on a two-tailed t-test (WT compared to rbl10-1 0.087 ³ p and rbl10-2 0.086³ p). Acyl compositions of total lipid extract (b), PG (c), DGDG (d), SQDG (e), and PC (f) are shown for wild type (WT) and the rbl10-1 and rbl10-2 mutant lines (n=3). Error bars report SE. Abbreviations of lipids are as defined in the legend of Figure 3. WT rbl11-1 rbl11-2 rbl11-3 Total (a) % l o M 50 40 30 20 10 (b) % l o M 60 50 40 30 20 10 WT rbl11-1 rbl11-2 rbl11-3 MGDG 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 0 0 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 Acyl Groups Figure 3.S2. Acyl composition of rbl11 total leaf lipids and MGDG. (a) Total leaf lipids and (b) MGDG (carbon # : double bond #), measured in three independent rbl11 alleles and WT, showing no change in 16:3 and 18:3 relative acyl chain abundance in the mutants compared to WT n=3. Error bars report SE. Acyl Groups 78 (a) i t h g e w h s e r f ¹ ˉ g • l o m p 160,000 120,000 80,000 40,000 (b) i t h g e w h s e r f ¹ ˉ g • l o m p (c) t i h g e w h s e r f ¹ ˉ g • l o m p 0 1600 1200 800 400 0 20000 16000 12000 8000 4000 0 JA WT rbl10-1 rbl10-2 0 JA-Ile 0 OPDA 20 60 20 60 0 20 Time (min) 60 Figure 3.S3. Changes in oxylipin levels in the rbl10 mutants. (a) LC-MS shows that rbl10-1 and rbl10-2 mutant lines accumulate significantly less jasmonic acid (JA) and (b) JA-Ile at 20 minutes after leaf wounding compared to wild type (WT, 0.006 ³ p, 0.004 ³ p and 0.044 ³ p, 0.064 ³ p respectively). (c) Both mutant lines accumulated less 12- Oxophytodienoic acid (OPDA) at 20 minutes after wounding than WT but not to a significant difference at the 0.05 level (0.081 ³ p, 0.090 ³ p respectively). Error bars represent SE, all panels (n=4); a two-tailed t-test was applied. 79 e u q i l i S / # d e e S 40 20 0 W T Figure 3.S4. Seed phenotypes of rbl10 plants. 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(2013) The phosphatidic acid binding site of the Arabidopsis Trigalactosyldiacylglycerol 4 (TGD4) protein required for lipid import into chloroplasts. J Biol Chem, 288, 4763-4771. Wang, Z. and Benning, C. (2011) Arabidopsis thaliana polar glycerolipid profiling by thin layer chromatography (TLC) coupled with gas-liquid chromatography (GLC). Journal of visualized experiments, 1-6. 85 Xu, C., Fan, J., Froehlich, J.E., Awai, K. and Benning, C. (2005) Mutation of the TGD1 chloroplast envelope protein affects phosphatidate metabolism in Arabidopsis. The Plant cell, 17, 3094-3110. Xu, C., Yu, B., Cornish, A.J., Froehlich, J.E. and Benning, C. (2006) Phosphatidylglycerol biosynthesis in chloroplasts of Arabidopsis mutants deficient in acyl-ACP glycerol-3-phosphate acyltransferase. Plant Journal, 47, 296-309. Yang, Y., Zienkiewicz, A., Lavell, A. and Benning, C. (2017) Coevolution of domain interactions in the chloroplast TGD1, 2, 3 lipid transfer complex specific to brassicaceae and poaceae plants. Plant Cell, 29. Yu, B., Wakao, S., Fan, J. and Benning, C. (2004) Loss of plastidic lysophosphatidic acid acyltransferase causes embryo-lethality in Arabidopsis. Plant Cell Physiol, 45, 503-510. 86 CHAPTER 4 RBL10 Protein Properties and Oligomeric State 87 Abstract Intramembrane proteolysis has been implicated in a variety of regulatory mechanisms in cells. A family of rhomboid proteases carry out proteolysis within the membrane and participate in numerous signaling pathways across nearly all organisms. While their roles are proving to be important to human health, characterization of their molecular mechanisms remains challenging. Though many rhomboid-like proteins are predicted in plants, no detailed molecular mechanism is available for any of them yet. Of the 13 predicted rhomboids in Arabidopsis thaliana, one rhomboid has been found to influence lipid metabolism in the chloroplast. Rhomboid-like protein 10 (RBL10) seems to alter the trafficking of phosphatidic acid (PA), a key lipid intermediate and signaling molecule, ultimately leading to a change in monogalactosyldiacylglycerol (MGDG) acyl composition in the thylakoid membranes. RBL10 also appears to be part of a large molecular weight complex greater than 660 kDa in size. Furthermore, RBL10, but not active site mutants, undergoes autolytic cleavage in its carboxy-terminal domain. This autocatalytic activity proves that RBL10 has protease activity, although other substrates are not yet known. The identity of the RBL10 complex components as well as the in vivo mechanism of autocleavage of RBL10 provide stepping stone towards a deeper understanding of how RBL10 participates in Arabidopsis lipid metabolism. Introduction Rhomboid family proteins carry out intramembrane proteolysis, utilizing a serine and histidine catalytic dyad to process peptides within the lipid bilayer of the membrane (Cho et al., 2019). Their roles vary depending on the organelle they associate with and the 88 tissues they are present in. Generally, members of the rhomboid family tend to carry out regulated intramembrane proteolysis, activating ligands (Lee et al., 2001, Urban et al., 2002), initiating peptide trafficking (Meissner et al., 2011), and enabling the oligomerization of complexes (Stevenson et al., 2007). Though they do not require an amino acid target consensus sequence for cleavage, they maintain selectivity while scanning the membrane for their substrates (Moin and Urban, 2012, Kreutzberger et al., 2019). Rhomboid proteases and their regulatory roles in mammals have gained medical relevance since the discovery of the family in Drosophila melanogaster (Bergbold and Lemberg, 2013, Freeman, 2016). While an appreciable number of rhomboid roles have been described so far, there are yet to be any rhomboids to be functionally characterized in plants even though many members are predicted in Arabidopsis thaliana, Populus trichocarpa, and Oryza sativa (Garcia-Lorenzo et al., 2006, Tripathi and Sowdhamini, 2006, Lemberg and Freeman, 2007). An active member of the rhomboid family has been confirmed with protease activity assays in Arabidopsis (Kanaoka et al., 2005) though the biological role still remains elusive. So far, the best studied plant rhomboid is found in the chloroplasts of Arabidopsis and is encoded by At1g25290 or the Rhomboid-like protein 10 (RBL10) gene. A number of publications probed the biological relevance of RBL10 as well as its mechanism, showing that knock-outs of RBL10 have altered pollen morphology, floral defects, and might influence jasmonic acid (JA) signaling (Knopf et al., 2012, Thompson et al., 2012). More recently, RBL10 has been implicated in Arabidopsis lipid metabolism and the utilization of chloroplast assembled phosphatidic acid (PA) for galactolipid biogenesis (Lavell et al., 2019). Though the involvement of RBL10 in lipid metabolism narrowed its potential biological function, the exact mechanism through which 89 this rhomboid-like protein affects PA metabolism or trafficking is not clear. This chapter focuses on the oligomeric state and proteolytic properties of RBL10 advancing our understanding of RBL10 function in plants. Results and Discussion The Oligomeric State of RBL10 Indicates Stable Protein Interactors An outstanding question surrounding rhomboid catalysis addresses the oligomeric state of rhomboids and probes the idea that they are obligate monomers. However, there are known rhomboid protein non-substrate interactors (Jeyaraju et al., 2006), but few examples of stable complexes involving rhomboids have been reported. Upon immunoblot analysis of RBL10-YFP-HA analyzed by separating solubilized chloroplast proteins under non-denaturing conditions using Blue-Native Polyacrylamide Gel Electrophoresis (BN-PAGE), it appears that RBL10 may be a part of a large molecular weight complex (Fig. 4.1). The observed band size is much larger than expected for a monomer or even dimer of the protein (~60 and ~120kDa respectively). The anti-HA signal shows three bands, one band nearing 1,000 kDa and the other two are ~400 and 450 kDa respectively. These findings were corroborated by using size exclusion chromatography (SEC), where solubilized chloroplast proteins of RBL10-YFP-HA- expressing Arabidopsis were separated on a Superdex 200 column. With larger MW proteins and complexes eluting earlier, we readily observed the elution of the full length RBL10-YFP-HA fusion protein in the earlier fractions corresponding to a MW of >660 kDa (Fig. 4.2). 90 The RBL10 containing complex could represent a new example of a protein complex involved in lipid metabolism in Arabidopsis. Currently, knowledge about the oligomeric states of lipid biosynthetic enzymes, or multiprotein complexes in which they play a possible role, is generally lacking. One complex involved in PA binding and lipid trafficking from the ER is an ABC transporter consisting of TRIGALACTOSYLDIACYLGLYCEROL 1, 2, and 3, (Xu et al., 2005, Awai et al., 2006, Lu et al., 2007, Lu and Benning, 2009, Roston et al., 2012, Yang et al., 2017)which likely interacts with two additional proteins, TGD4 and 5 (Xu et al., 2008, Wang et al., 2012, Fan et al., 2015).Though the TGD complex is also of a large molecular weight, the RBL10 protein does not seem to overlap with the TGD complex using different sizing approaches. As a next step, we characterized the protein interactome of RBL10 to gain insight into its molecular function. Furthermore, it is not known at this time, what the endogenous protein substrate of RBL10 is, and it seemed possible that it might be part of the complex. Co-Immunoprecipitation of RBL10 Yields Putative Protein Interactors To identify possible protein interactors and substrates of RBL10, we used a co- immunoprecipitation approach. The RBL10-YFP-HA fusion protein and with its associated proteins present in chloroplast extracts from transgenic plants expressing the RBL10-YFP-HA construct in the rbl10-1 mutant background were bound to magnetic beads with an antibody specific for the HA-tag present on RBL10-YFP-HA. To control for non-specific binding, an isotype control antibody was used, which has no specificity toward HA or other proteins present in the chloroplast lysate. Both samples, were submitted for mass spectrometry to identify any proteins immunoprecipitated with each 91 set of beads. A few hundred proteins were detected in both sample sets. However, upon comparing the HA-specific and control samples and across three experimental replicates, a small subset of recurring proteins was identified specific to the HA-specific sample (Table 4.1). There was a total of 22 proteins co-immunoprecipitated with RBL10-YFP-HA in at least 2 of the 3 replicates. Of those 22, a single protein (#15) appeared in all three experimental replicates, annotated as ribosomal protein S1 (RPS1) with gene ID At5g30510 (Table 4.1). There is some evidence that RPS1 does indeed function as a ribosomal protein in the chloroplasts of Arabidopsis. It recognizes mRNAs and is important in the translation of RNAs encoding thylakoid membrane proteins (Romani et al., 2012). There is additional evidence that RPS1 might facilitate retrograde signaling from the chloroplast in response to heat stress, and that RPS1 function is important for normal thylakoid ultrastructure (Yu et al., 2012). In light of potential physical interactions between RPS1 and RBL10, the mechanistic relationship between changes in PA metabolism and heat stress response is not immediately clear. However, it is possible that heat-induced stress could require an RBL10-mediated adjustment of the thylakoid membrane lipid composition through varying the accessibility of different PA pools, which affects the acyl chain length of the most abundant thylakoid lipid, MGDG. However, RPS1 does not seem to be associated with any membrane, a prerequisite for a rhomboid substrate. While it is unlikely that RPS1 is the direct protein substrate of RBL10, it is plausible that it could be stably interacting with a soluble region of RBL10, such as its amino terminus, and serve a regulatory role. The remaining 21 proteins which appear in 2 of 3 experimental replicates have annotations related to processes such as photosynthesis, protein import, chaperonin 92 function, carbohydrate metabolism, signaling, as well as one protein possibly related to fatty acid metabolism (Table 4.1). Before any more conclusions can be drawn from any of these proteins as well as RPS1, their physical interactions with RBL10 must be tested in different ways. This can be accomplished through a split-ubiquitin yeast two-hybrid approach (Obrdlik et al., 2004). Once there is additional evidence obtained about the direct interaction with RBL10, or interactions between any of these individual components, further conclusions can be drawn about the molecular function of these candidate proteins in the RBL10 complex. RBL10 is Autolytic Towards its Carboxy-Terminal Domain Protease activity assays have been challenging to perform with purified RBL10 which was labeled at its carboxy-terminal domain (CTD). Purification of the WT protein appeared unsuccessful, where yields were always quite low, and it seemed implausible that so little protein was being produced in Nicotiana benthamiana as well as Escherichia coli. Initially this was presumed to due to the deleterious effects of producing an active protease with many hydrophobic domains. However, there was suspicion that the CTD may be subjected to proteolytic cleavage by the action of RBL10 or another protease. To further investigate, the predicted catalytic residues of RBL10, Serine 240 and Histidine 293, which were previously identified (Chapter 3) were mutagenized using PCR to convert both residues to alanine, resulting in RBL10S240A and RBL10S240A;H293A variants. The expression of these mutant constructs in E. coli, yielded a CTD glutathione-s-transferase (GST)-tagged RBL10 protein (Fig. 4.3). The WT RBL10-GST protein cannot be detected with antibody raised against GST, however, the catalytically inactive proteins are detected 93 with the anti-GST antibody. This finding suggested that the CTD of RBL10 is only cleaved when the catalytic residues are present and that RBL10 cleaves its own CTD. Since there are six main transmembrane segments in the conserved rhomboid fold, the prediction of a seventh transmembrane domain at the CTD may indicate that it is this segment of the protein which gets cleaved by RBL10. While a CTD tag is not detectable on WT RBL10, an amino-terminal domain (NTD) tagged RBL10 yielded detectable protein production in E. coli even for WT RBL10 around 70 kDa in size (Fig. 4.4). Autolytic cleavage has previously been reported for the human PARL, where the cleavage product contains a nuclear targeting sequence, which is conserved in mammals, and travels to the nucleus upon release from the mitochondrion (Sik et al., 2004). Though we can observe the cleavage of RBL10 CTD, we do not yet know the fate of the released peptide. Determining the exact site of cleavage as well as any secondary localizations of the released fragment could give us greater insight into the function of this autolysis. While autolysis appears to happen as protein is produced in a bacterial host, the ability to detect an intact fusion as well as a retained CTD fragment in isolated chloroplasts might give some clues about the coordination of this autocleavage. Interestingly, we were also able to detect a smaller RBL10CTD-YFP-HA in the later fractions of the SEC corresponding to a monomeric form of the fragment (Fig. 4.2). It is unclear whether the remaining amino terminal portion of the protease remains with the complex. Since RBL10 appears to have both a full fusion and the cleaved fragment, while in bacterial cells the autocatalysis occurs as soon as protein is made, it is possible that the interaction with complex members can control autocatalysis of RBL10. The CTD fragment is only found in a monomeric state in the SEC analysis (Fig. 4.2), perhaps indicating that dissociation 94 from the complex and autocatalysis coincide. It is not currently clear, whether the dissociation from the complex induces autocatalysis or the cleavage of the CTD causes dissociation from the complex. Since we don’t yet have an antibody against RBL10 protein, we cannot determine whether the processed RBL10 form remains complex bound or if it dissociates altogether. Understanding the relationship between autocatalysis and complex residency of RBL10 is an important next step towards solving its molecular mechanism. Catalytic Mutant Constructs of RBL10 Cannot be Stably Transformed into the rbl10- 1 Mutant Background The causal relationship between the observed lipid phenotype in the rbl10 mutants and the RBL10 gene was shown by rescuing the mutant line with a WT copy of RBL10 (Lavell et al., 2019) (See Chapter 3). It was then important to establish whether the proteolytic function of RBL10 was important to its role in lipid homeostasis. The point mutants which were generated using PCR mutagenesis, RBL10S240A and RBL10S240A;H293A were cloned into the same pEARLEYGATE101 plant expression vector as used for the WT sequence, possessing a BASTA herbicide marker for screening transformants. However, unlike the WT construct, the floral dipping transformation of constructs bearing the point mutants of RBL10 with Agrobacterium tumefaciens did not yield any transformants. With two attempted transformations and thousands of seeds screened for BASTA resistance, no stable transgenic lines were recovered. This indicated that there is possible embryo lethality when introducing non-catalytic RBL10 mutant protein in the rbl10-1 and WT background, with the mutant construct behaving like a dominant negative mutation. A 95 conclusion that could be drawn from this finding, is that RBL10 is part of an essential complex that is necessary for the function of the plastid galactolipid pathway. We know that this pathway is not essential for embryogenesis and plant growth from the knock out mutants of the ATS1 gene (Kunst et al., 1988). However, introducing a non-active RBL10 protein seems to imply that the presence of RBL10 protein itself might be influencing the functionality of the complex. In fact, RBL10 is part of a large molecular weight complex as we have shown above, although we do yet not know for certain its components in the wild type or in the rbl10 mutant. With its autolytic activity demonstrated in this study, perhaps RBL10 must process its CTD in order to dissociate from the complex and turn over. Without the ability to process its own CTD, the complex cannot turn over resulting in deleterious effects on developmental processes. It is possible that there is a regulatory step involving this complex which determines the flux through the two lipid biosynthetic pathways. Without any RBL10 protein present, the complex may favor the ER pathway which is essential (Xu et al., 2005), and with a copy of RBL10 protein present, the complex favors the plastid pathway. However, if a null copy is introduced and having proteolytic ability is essential for RBL10 to dissociate from the complex, the null protein would prevent switching between the two biosynthetic pathways. If this is the case, then a null copy of RBL10 would in fact cause embryo lethality due to the plant’s inability to utilize the essential ER pathway during embryogenesis. Due to our lack of understanding of the exact mechanism of RBL10 complex formation and the role of its autocatalytic activity in complex association/dissociation, we cannot fully explain why stable transgenic lines expressing null copies of RBL10 have not been successfully generated. It would be instructive to test whether the non-functional 96 versions of RBL10 can be successfully produced with transient expression in Nicotiana benthamiana to then test the effects on lipid composition and RBL10 complex formation. It is possible that autocatalysis of RBL10 is important for dissociation from the complex, and without being able to do so, there are deleterious metabolic consequences for the plant. Materials and Methods Plant Growing Conditions and Chloroplast Isolation Arabidopsis seeds were sterilized (1 part bleach and 2 parts water solution with 0.1% Triton X-100) and sown on MS media supplemented with 1% sucrose. Approximately, 30 mg of seeds per plate were evenly distributed. Plates were sealed with 3M micropore tape, stratified for 3 days at 4°C, then placed into a 100 µmol light growth, 22°C, with a 16 hr day and 8 hr night environment in a Percival chamber. Tissue was harvested for chloroplast isolation between 3 and 4 weeks of age. Plant lines were used as noted in each experiment including Col-0 (WT), rbl10-1, and the rbl10-1 complemented line expressing RBL10-YFP-HA under 35S Cauliflower Mosaic Virus (CaMV) promoter using the pEARLEYGATE101 vector. Chloroplasts were isolated as previously described (Lavell et al., 2019). SDS-PAGE and Immunoblot Analysis Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) was carried out using Bio-Rad precast 10-12% acrylamide gels (https://www.bio-rad.com), or hand-cast 10-12% acrylamide gels. Samples were boiled in Laemmli buffer (50mM Tris-HCL pH6.8, 97 2% SDS, 0.1% Bromophenolblue, 10% glycerol at 1x) at 95°C for 10 mins, centrifuged for 3 minutes at 13,000 rpm, and loaded onto the gel. A protein standard mixture was used for size reference (Bio-Rad Precision Plus Dual Stained Molecular Weight marker (https://www.bio-rad.com)). For running buffer, Tris-Glycine (25 mM Tris, 192 mM Glycine) containing 0.1% sodium dodecyl sulfate SDS was used. Proteins were separated at ~150V until the loading dye front was just running off the bottom of the gel. Resulting gels were either directly stained with a Coomassie staining solution (1.25g Coomassie brilliant blue, 400mL methanol, 100mL acetic acid, 500mL dH2O) or were used for transferring to a PVDF membrane. Immunoblot transfer was performed using the wet transfer method, using a Tris-Glycine buffer (25 mM Tris, 192mM Glycine, 20% methanol). Protein transfer was achieved with 100 V over 1.5 h chilled on ice with stirring. A 5% milk solution in TBS-T (100 mM Tris, 155 mM NaCl, 0.05% Tween 20) was used to block the resulting blots for an hour at RT or overnight at 4°C. After blocking, blots were incubated with the antibody specified in a 5% milk-TBS-T solution for 1 h at room temperature or overnight at 4°C. Washes in between primary and secondary antibodies were performed with TBS-T buffer, washing the blot three times for 5-15 mins each time. Chemiluminescence was used to detect presence of protein of interest. Blue Native PAGE Arabidopsis isolated chloroplasts were freshly prepared on the day of use, using WT, rbl10-1, and rbl10-1 complemented line expressing RBL10-YFP-HA under 35S promoter. Chloroplasts were pelleted at 700 x g, then proteins were solubilized in the BN-PAGE protein solubilization buffer consisting of 333 µL of 3x Gel Buffer (150 mM Bis-tris/HCL, 98 1.5 M 6-aminocapronic acid, pH 7.0), 125 µL of 80% glycerol, 100 µL of 10% n-dodecyl b-D-maltoside (DDM), 10 µL of 100mM phenylmethylsulfonyl fluoride (PMSF), and water to 1 mL. The final concentration was 1 mg/mL chlorophyll equivalents and the solution was allowed to incubate on ice for 30 mins with occasional tapping to mix. The resulting chloroplast lysate was clarified with a 10 min centrifugation at 100,000 x g. The supernatant was transferred to a clean tube and 5% SERVA Blue G (SERVA No. 35050) dye was added to the solution for a final concentration of 3.0%. An unstained native gel marker was used for size reference (NativeMark; www.ThermoFisher.com). Chloroplast protein complexes were separated on a 4-16% Bis-Tris polyacrylamide gel overnight at 30V in 4°C. The resulting gel was either stained directly using Coomassie staining solution or transferred to a PVDF membrane for Immunoblot analysis. To transfer the proteins to PVDF, the gel was first boiled in a boiling buffer solution (3.3% w/v SDS, 4% 2-mercaptoethanol, and 65mM Tris buffer) for 30 min at 80°C using a glass petri dish on a hot plate in a fume hood with gentle mixing. The gel was then washed in transfer buffer to remove excess SDS and reducing agent. Proteins were transferred as described above. The resulting PVDF membrane was fixed in 8% acetic acid. Any transferred dye was removed with methanol, followed by washing with water. Immunoblot analysis proceeded as described above. RBL10-YFP-HA was detected using an anti-HA antibody from rat directly conjugated to horseradish peroxidase (HRP) enzyme at a 1:1000 dilution (3F10, Roche/Sigma Aldrich, www.sigmaaldrich.com). 99 Size Exclusion Chromatography A standard protein mixture was purchased from Sigma-Aldrich (Catalog# 69385), including thyroglobulin (670k Da), bovine g-globulins (150 kDa), chicken egg albumin (44.3 kDa), bovine ribonuclease A type I-A (13.7 kDa), and p-aminobenzoic acid (137 Da). The standard protein mixture was supplied as a lyophilized powder corresponding to 30 mg of protein. The powder was resuspended with 1 mL of deionized water and any undissolved solids were removed with centrifugation. A GE Superdex 200 (www.gelifesciences.com) column was equilibrated with phosphate buffered saline (PBS) pH 7.4 with 0.01% DDM at 4°C. A 100 µL injection loop was used to load 100 µL of the standard mixture onto the equilibrated column. Proteins were separated by size exclusion chromatography using a GE AKTA purifier system (www.gelifesciences.com) at 0.05 mL/min. Fractions of 0.5 mL were collected from the time of injection. A total of 72 fractions were collected, equal to 1.5 times column volume. Protein elution was monitored by absorbance at 280 nm, signals corresponding to individual proteins in the standard mixture were observed for reference. Arabidopsis isolated chloroplasts were freshly prepared the day of use, using the rbl10-1 complemented line expressing RBL10-YFP- HA. Chloroplasts were pelleted at 700 x g, then proteins were solubilized in the BN-PAGE protein solubilization buffer as described above to a 1 mg/mL chlorophyll equivalent concentration, allowing the solution to incubate on ice for 30 mins with occasional tapping to mix. The resulting chloroplast lysate was clarified with a 10 min centrifugation at 100,000 x g. The supernatant was removed and a 100 µL, equal to 100 µg chlorophyll equivalents, was loaded onto a Superdex 200 SEC column equilibrated with PBS pH 7.4 with 0.01% b-dodecyl-n-maltoside (DDM). The chloroplast lysate was separated at 0.05 100 mL/min flow rate using the same method as the standard mixture. Again, A280 was used to monitor protein elution off the column. Standard proteins were confirmed with SDS- PAGE of the fractions corresponding to the A280 signals. Elution of RBL10-YFP-HA was observed by immunoblot analysis of the resulting fractions in the same range as the A280 signals for the standards, using an anti-HA antibody from rat directly conjugated to horseradish peroxidase (HRP) enzyme at a 1:1000 dilution (3F10, Roche/Sigma Aldrich, www.sigmaaldrich.com). Co-Immunoprecipitation Magnetic Dynabeads with Protein G were prepared prior to the experiment. Mouse IgG specific for hemagglutinin (HA) (Biolegend 16b32) and a mouse K-isotype control IgG were used to couple to the Protein G Dynabeads, using 7.5 µg per 50 µL of beads. Chloroplast protein lysate was prepared as described above for BN-PAGE without the addition of SERVA Blue G dye. Each set of IgG coupled Dynabeads was incubated with the chloroplast protein lysate at room temperature with end over end mixing for 1 hr. Beads were collected with a magnet, supernatant removed, and the beads were washed three times with 1x PBS pH 7.4, DDM 0.02%. The resulting beads were submitted for mass spectrometry identification of peptides present. Samples were washed with ice-cold 50mM ammonium bicarbonate, 3x and finally resuspended in 10uL of 5ng/µL trypsin in the same buffer. Beads were incubated at 37°C for 5hrs to digest proteins. Supernatant was removed following centrifugation, placed into a new tube and the peptide solution acidified to 3% trifluoroacetic acid (TFA). Samples were then purified using c18 stage-tips (Shevchenko et al., 1996) and dried by vacuum 101 centrifugation. Peptides were then re-suspended in 2% acetonitrile/0.1%TFA to 20µL. From this, 10µL were automatically injected by EASYnLC 1200 (www.thermo.com) onto a Thermo Acclaim PepMap RSLC 0.1mm x 20mm C18 trapping column and washed with Buffer A for ~5min. Bound peptides were eluted onto a Thermo Acclaim PepMap RSLC 0.075mm x 500mm C18 analytical column over 35min with a gradient of 8%B to 40%B in 24min, ramping to 90%B at 25min and held at 90%B for the duration of the run (Buffer A = 99.9% Water/0.1% Formic Acid, Buffer B = 80% Acetonitrile/0.1% Formic Acid/19.9% Water). Column temperature was maintained at 50C using an integrated column heater (PRSO-V2, www.sonation.com). Eluted peptides were sprayed into a ThermoFisher Q-Exactive HF-X mass spectrometer (www.thermo.com) using a FlexSpray spray ion source. Survey scans were taken in the Orbi trap (60000 resolution, determined at m/z 200) and the top 10 ions in each survey scan are then subjected to automatic higher energy collision induced dissociation (HCD) with fragment spectra acquired at 15,000 resolution. The resulting MS/MS spectra are converted to peak lists using Mascot Distiller, v2.6.1 (www.matrixscience.com) and searched against all entries in the TAIR, v10, protein sequence database (downloaded from The Arabidopsis Information Network, (www.arabidopsis.org)), appended with common laboratory contaminants (downloaded from www.thegpm.org, cRAP project), using the Mascot searching algorithm, v 2.6. The Mascot output was then analyzed using Scaffold Q+S, v4.9.0 (www.proteomesoftware.com) to probabilistically validate protein identifications. Assignments validated using the Scaffold 1%FDR confidence filter are considered true. 102 Proteins unique to the RBL10-YFP-HA sample or twice as abundant compared to the control samples in at least 2 of the 3 experimental replicates were considered putative protein interactors. The protein names, gene IDs, molecular weight, net enrichment as well as the occurrence throughout experimental replicates is shown in Table 4.1 . Protein production in E. coli. Four strains bearing pDEST24 gateway vector (www.ThermoFisher.com) containing the following were generated: a 50 amino acid peptide of YFP (EV), RBL10 WT gene, RBL10S240A single serine mutant gene, and RBL10S240A;H293A serine/histidine double mutant gene. The constructs were generated by recombining the listed genes in pENTR/SD/D/Topo vector (www.ThermoFischer.com) with the pDEST24 vector using LR Clonase II (www.ThermoFischer.com). Starter cultures of BL21(DE3) cells transformed with the four pDEST24 constructs were grown overnight from glycerol stock in Luria broth (LB) with ampicillin at 37°C. The following day, 25 mL cultures were inoculated in flasks at a cell density corresponding to an OD600 of 0.1, and grown with shaking at 37°C until the cell density was 0.4-0.7. An initial sample was taken, a volume equal to an OD600 of 0.5, at timepoint 0, the cells were spun down, supernatant removed, and snap frozen in liquid nitrogen, then kept at -80°C. Expression of plasmids was induced by the addition of 20 µM Isopropyl β-d-1-thiogalactopyranoside (IPTG) to each flask. Samples were taken at later times with the same OD600 across all timepoints. Once all the samples were collected, the pellets were resuspended in 40 µL of water and 10 µL of 5 x Laemeli Buffer was added, the samples were boiled at 95°C for 10 mins, then 20 µL were loaded onto a acrylamide gel and separated as described above followed by immunoblot analysis. To 103 probe for the presence of RBL10 protein fusions with GST, a rabbit anti-GST primary antibody was used at a 1:5000 dilution. For production of NTD tagged maltose-binding protein (MBP)-RBL10 fusion, the pDEST-His-MBP vector was obtained from Addgene (www.addgene.com). WT and single serine 240 RBL10 constructs in pENTR/SD/D-topo were recombined as described above with the pDEST-His-MBP vector. The generated constructs were transformed into BL21(DE3) cells. Cultures of the resulting strains containing WT RBL10 and RBL10S240A in the pDEST-His-MBP vector were grown up at 37°C overnight in LB with ampicillin. The following day, 25mL cultures of LB with ampicillin were inoculated from the overnight cultures to an OD600 of 0.1. The cultures were then grown at 37°C until OD600 of 0.6, and were induced with 20 µM IPTG, the temperature was reduced to 16°C. After overnight growth, the cultures were harvested by centrifugation. Pellets were resuspended in 10mL of lysis buffer (PBS pH7.4, 10mM MgCl, 8mg/mL lysozyme, 1 tablet of EDTA-free mini complete protease inhibitor (Roche, www.sigmaadrich.com)). The cells were disrupted by sonication (1 sec on, 1 sec off, for 3 minutes total) on ice. Cellular debris were spun down at 24,000 x g for 12 minutes. Membranes were pelleted at 100,000 x g for 1 hr. The membrane fraction was resuspended in 2.5mL of lysis buffer amended with 0.2% TritonX- 100. The solubilized membranes were passed three times over 2mL of amylose resin prepared by the manufacturer’s recommendation (www.NEB.com), Wash and elution steps were performed as per manufacturer’s recommendation. The eluted fractions were desalted into PBS pH 7.4, using an amicon centrifugal unit (Millipore) with a molecular weight cutoff of 30kDa. Protein concentration was determined using BCA (Pierce, www.thermofisher.com), and 20ug of protein from both RBL10 WT sample and 104 RBL10S240A were prepared for SDS-PAGE as described above and loaded onto a 10% polyacrylamide gel twice, half the gel was used for Western blot transfer and the other half was used for direct staining with Coomassie. For Western blot analysis anti-MBP antibody from rabbit was used at a 1:75,000 dilution. Acknowledgement This work was possible due to the help and generosity of many individuals. For performing the size exclusion chromatography, Dr. Curtis Wilkerson graciously allowed the use of his instrument and column as well as provided valuable insight. Dr. Mingzhu Fan provided careful training on the use of the Akta instrument and assisted with operation. Cameron De La Mora assisted on generating Figure 4.3. For critical discussions on co- immunoprecipitation experiments and BN-PAGE, Tomomi Takeuchi deserves a big thank you. Identification of potential protein interactors in Table 4.1 would not have been possible without Doug Whitten at the MSU Proteomics Core. Financial support for this work was primarily from the Division of Chemical Sciences, Geosciences and Biosciences, Office of Basic Energy Sciences of the United States Department of Energy (Grant DE-FG02-91ER20021). In addition, Anastasiya Lavell was partially supported by a fellowship from Michigan State University under the Training Program in Plant Biotechnology for Health and Sustainability (T32-GM110523). 105 APPENDIX 106 binding protein 3 chloroplasts 159 dehydrogenase Gene Annotation bacterial/chloroplast # 1 PGK1 - phosphoglycerate kinase 1 2 ATPC1 - ATPase, F1 complex, gamma subunit protein 3 Transketolase 4 LHCB3 - light-harvesting chlorophyll B- 5 PSB27 - photosystem II family protein 6 ATPase, F0 complex, subunit B/B', 7 ZKT - protein containing PDZ domain, a K- box domain, and a TPR region 8 APX4 - ascorbate peroxidase 4 9 TOC159 - translocon at the OEM of 10 CPN60B, LEN1 - chaperonin 60 beta 11 EDA9 - D-3-phosphoglycerate 12 SBPASE - sedoheptulose-bisphosphatase 13 CLPC1 - CLPC homologue 1 14 ATPB - ATP synthase subunit beta 15 RPS1 - ribosomal protein S1 16 RPL2.1 - ribosomal protein L2 17 Pyridine nucleotide-disulphide oxidoreductase family protein 18 Thioredoxin superfamily protein 19 PRK - phosphoribulokinase 20 FTSH1 - FTSH protease 1 21 ACP4 - acyl carrier protein 4 22 CPN20 - chaperonin 20 Accession Number AT3G12780 AT4G04640 AT3G60750 AT5G54270 AT1G03600 AT4G32260 MW kDa 50 41 80 29 19 24 37 AT1G55480 AT4G09010 38 AT4G02510 161 AT1G55490 64 63 AT4G34200 AT3G55800 42 20 AT5G50920 54 ATCG00480 45 AT5G30510 ATCG00830 30 52 AT1G74470 AT3G11630 29 44 AT1G32060 77 AT1G50250 AT4G25050 15 27 AT5G20720 Net Enrichment Occurrence 10.483 4.765 4.765 4.765 3.7131 2.6612 2.859 1.8071 1.9833 1.906 1.906 1.906 1.906 0.99167 0.99167 0.99167 0.99167 0.95299 0.95299 0.95299 0.95299 0.95299 2 2 2 2 2 2 2 2 2 2 2 2 2 2 3 2 2 2 2 2 2 2 Table 4.1. Co-Immunoprecipitation with RBL10-YFP-HA using isolated chloroplast protein lysate, results in recurring appearance of select proteins. Proteins that were identified as at least twice as abundant in the anti-HA sample versus control are described here with their gene annotation, their gene ID, predicted molecular mass, the enrichment in the anti-HA sample compared to the control, and the recurrence out of three experimental replicates. 107 a b c d kDa 1,000 480 kDa kDa 1,000 480 kDa WT rbl10-1 rbl10-1:RBL10-YFP-HA rbl10-1 complemented lines producing RBL10-YFP-HA protein. Figure 4.1. Blue Native PAGE of solubilized chloroplasts from WT, rbl10-1 mutant, and (a) Immunoblotting of the resulting BN-PAGE gel, blotted against HA, shows only signal in the complement line with a band above 1,000 kDa and (b) two bands below 480 kDa. (c) The immunoblot stained with Coomassie showing the region corresponding to 1,000 kDa and (d) 480kDa. For each genotype 10, 20, and 30 µL of chloroplast prep were loaded. 108 Figure 4.2. Size Exclusion Chromatography of protein standards and solubilized chloroplasts of rbl10-1:RBL10-YFP-HA line show that RBL10 migrates in a complex at a high MW. (a) The standards used were: 1, Thyroglobulin (660 kDa); 2, Bovine g- globulins (150 kDa); 3, Chicken albumin (44.3 kDa); 4, Ribonuclease A type I-A (13.7 kDa); 5, P-aminobenzoic acid (pABA)(137Da). They are resolved on a Superdex 200 column with A280 monitoring of protein elution over time. (b) Elution of solubilized chloroplast proteins of the rbl10-1:RBL10-YFP-HA line from the Superdex 200 column, monitored with A280 over time. (c) SDS-PAGE of the fractions containing separated protein standards from (a), with the exception of pABA, show each standard and the respective fraction number. (d) Immunoblot analysis of fractions containing separated chloroplast proteins from (b), detecting signal with HA antibody. 109 Figure 4.3. Production of C-Terminally tagged RBL10-GST in E. coli. The absence of the fusion protein is observed for WT constructs, and this is rescued by mutagenesis of the active site serine and histidine residues. (a) The fusion protein of RBL10-GST (~64kDa) can be detected after 1 and 3 h post IPTG induction by anti-GST antibody for the single and double catalytic mutant (RBL10S240A-GST and RBL10S240A:H293A-GST) but not WT RBL10-GST, using immunoblot analysis. Equal sample loading was ensured by normalizing to OD600 when samples were taken at each time point. (b) Coomassie-Blue stained blot of the PVDF membrane above in (a). 110 a RBL10 WT RBL10 S240A b RBL10 WT RBL10 S240A kDa 130 100 70 55 40 35 25 15 kDa 130 100 70 55 40 35 25 15 Figure 4.4. The fusion protein of MBP-RBL10 WT and single catalytic mutant can be detected with anti-MBP antibody. (a) The WT fusion protein MBP-RBL10 appears on anti-MBP Immunoblot when the tag is located on the amino-terminal domain of RBL10. 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PLoS Genet, 8, e1002669. 115 CHAPTER 5 Arabidopsis DGD1 SUPPRESSOR 1 is a Subunit of the Mitochondrial Contact Site and Cristae Organizing System and Affects Mitochondrial Biogenesis is published further edited and in The Plant Cell This work has been (http://www.plantcell.org/content/31/8/1856). Anastasiya Lavell contributed to this work by performing polar lipid profiling of total leaf tissues as well as isolated chloroplasts and mitochondria of the lines used in this study. She analyzed and interpreted the data and assisted in the writing of the sections discussing lipids. Li, L., Lavell, A., Meng, X., Berkowitz, O., Selinski, J., van de Meene, A., Carrie, C., Benning, C., Whelan, J., De Clercq, I. and Wang, Y. (2019) Arabidopsis DGD1 SUPPRESSOR 1 is a subunit of the Mitochondrial Contact Site and Cristae Organizing System and affects mitochondrial biogenesis. Plant Cell. 116 Abstract Mitochondrial and plastid biogenesis require the biosynthesis and assembly of proteins, nucleic acids and lipids. In Arabidopsis thaliana the mitochondrial outer membrane protein DGD1 SUPPRESSOR 1 (DGS1) is part of a large multi-subunit protein complex, which contains the mitochondrial contact site and cristae organizing system (MICOS) 60 kDa subunit (MIC60), the translocase of outer mitochondrial membrane 40 kDa subunit (TOM40), the translocase of outer mitochondrial membrane 20 kDa subunits (TOM20s) and the Rieske FeS protein (RISP). A point mutation in DGS1 (dgs1-1), altered the stability and protease accessibility of this complex. This altered mitochondrial biogenesis, mitochondrial size, lipid content and composition, protein import and respiratory capacity. Whole plant physiology was affected in the dgs1-1 mutant as evidenced by tolerance to imposed drought stress and altered transcriptional responses of markers of mitochondrial retrograde signaling. Putative orthologs of Arabidopsis DGS1 are conserved in eukaryotes, including the NCA2 (Nuclear Control of ATP Synthase 2) protein in yeast (Saccharomyces cerevisiae), but lost in Metazoa. The genes encoding DGS1 and NCA2 are part of a similar co-expression network including genes encoding proteins involved in mitochondrial fission, morphology and lipid homeostasis. Thus, DGS1 links mitochondrial protein and lipid import with cellular lipid homeostasis and whole plant stress responses. Introduction Mitochondrial biogenesis requires the import of proteins, RNAs and lipids (Schneider, 2011, Mesmin, 2016, Wiedemann and Pfanner, 2017). The signals and machinery for the import and assembly of proteins are well studied, and one emerging 117 theme is that many proteins are dually targeted to both mitochondria and chloroplasts, highlighting the coordination of function of these organelles (Murcha et al., 2014). Both mitochondria and plastids contain their own genome, but rely on the import of nuclear- encoded proteins for the replication and expression of their genomes, with many of the involved proteins dually targeted (Elo et al., 2003, Carrie et al., 2009). While some lipids such as cardiolipin are produced in the mitochondria themselves, mitochondrial lipid biogenesis also depends on the import of lipids from other sites of synthesis, i.e., the endoplasmic reticulum and plastids (Michaud et al., 2017). Lipid transfer/import into mitochondria takes place via contact sites and is non-vesicular in nature. However, little is known about the proteins involved in this process in plants. In contrast, in yeast (Saccharomyces cerevisiae) many of the proteins involved have been identified and characterized (Michaud et al., 2017). In yeast and mammals, the mitochondrial contact site and cristae organizing system (MICOS/MINOS/MitOS), which bridges the inner mitochondrial membrane (IMM) and the outer mitochondrial membrane (OMM), has multi-faceted functions in mitochondrial morphology, protein import and abundance, oxidative phosphorylation as well as lipid biogenesis and content (van der Laan et al., 2016, Schorr and van der Laan, 2018). The components of MICOS are conserved in yeast and humans, with at least six subunits in yeast and nine subunits in humans (van der Laan et al., 2012, Kozjak-Pavlovic, 2017). A phylogenetic analysis for eukaryotes proposed that only two core components of MICOS, MIC60 and MIC10, were conserved in plants (Munoz-Gomez et al., 2015a, Munoz- Gomez et al., 2015b). A study in Arabidopsis revealed that MIC60, interacting with the translocase of the outer membrane (TOM) via TOM40, forms part of a mitochondrial 118 transmembrane lipoprotein complex (MTL), and affects mitochondrial lipid trafficking (Michaud et al., 2016). Mitochondrial membrane lipid homeostasis, including biosynthesis and transfer of phospholipids and cardiolipin, is controlled by MICOS-mediated membrane contact sites with the endoplasmic reticulum (ER) (Aaltonen et al., 2016, Wideman and Munoz-Gomez, 2016). In plants, transfer of lipids from plastids to mitochondria has also been detected and is also dependent on physical contact sites (Jouhet et al., 2004). During phosphate (Pi) limitation, phospholipids are replaced by non-phosphorous galactoglycerolipids synthesized in plastids (Härtel et al., 2000), although some galactoglycerolipids are also found in mitochondria under non-limiting phosphate growth conditions (Jouhet et al., 2004, Michaud et al., 2016). The regulatory mechanism behind determining the amount of mitochondrial galactoglycerolipids are unknown, as are the components responsible for their transfer from plastids to mitochondria. The bulk of the galactoglycerolipids, i.e., mono- and digalactosyldiacylglycerol (MGDG and DGDG), are synthesized at the inner and outer envelope membranes of the plastid, respectively, and they are highly abundant in the thylakoid membranes of the chloroplast. Galactolipids are unique to photosynthetic membranes under normal conditions. However, in response to Pi deprivation, galactolipids are exported to extra- plastidic membranes (Moellering and Benning, 2011). Galactolipid biosynthesis is carried out in the chloroplast envelope membranes by a set of MGDG synthases (MGDs) that catalyze the formation of MGDG followed by DGD synthases (DGDs) that use MGDG to form DGDG (Benning and Ohta, 2005). MGD1 and DGD1 are constitutively expressed while MGD2/3 and DGD2 are induced in response to phosphate limitation (Awai et al., 119 2001, Kelly and Dormann, 2002). To identify regulators of galactoglycerolipid biosynthesis in response to Pi deprivation, a genetic suppressor screen was conducted in the Arabidopsis thaliana dgd1 mutant background. A dgd1 suppressor mutant allele (dgd1 suppressor 1, dgs1-1) was identified that constitutively activated the DGD1-independent pathway for DGDG biosynthesis (Xu et al., 2003, Xu et al., 2008). The affected protein, DGS1, has two membrane-spanning domains and was localized to the OMM as part of a larger protein complex (Xu et al., 2008, Moellering and Benning, 2010). It contains a conserved domain also found in a yeast protein identified in a genetic screen for assembly of the ATP synthase complex, Nuclear Control of ATP Synthase 2 (NCA2) (Camougrand et al., 1995). In this study, we demonstrated that the OMM protein DGS1 is a novel subunit of MICOS in plants. We showed that DGS1 constituted part of a multi-subunit protein complex with MIC60. A point mutation of DGS1 affected chloroplast and mitochondrial lipid content and composition as well as mitochondrial biogenesis and function, which were also the effects of MICOS in yeast and mammals (van der Laan et al., 2016, Schorr and van der Laan, 2018). DGS1 was conserved across eukaryotes but absent from Metazoa and Alvaeolata. NCA2, the yeast putative ortholog of DGS1, and DGS1 had a similar co-expression network including proteins involved in mitochondrial biogenesis. 120 Results DGS1 is present in a multi-subunit protein complex with MIC60, TOM40, TOM20s and RISP Using immunoblotting following blue native PAGE (BN-PAGE), immunoprecipitation and cross-linking with the membrane-permeable chemical crosslinker disuccinimidyl glutarate (DSG), DGS1 was found to be present in a multi-subunit protein complex that contains MIC60, TOM40, TOM20s and the Rieske FeS protein (RISP) of the cytochrome bc1 complex (Figure 5.1). Mass spectrometry of excised bands revealed that DGS1 co- migrated with MIC60 in the oxidative phosphorylation complexes (OXPHOS), complex V, complex III and complex F1 (Figure 5.1A and Supplemental Dataset 1). The immunoblot following BN-PAGE revealed the majority of DGS1 was detected in complex III (Figure 5.11A), while MIC60 co-migrated with a variety of respiratory complexes (Figure 5.1A). Immunoprecipitation using a DGS1 antibody pulled down MIC60, TOM40, TOM20-2 and RISP, while the MIC60 antibody pulled down DGS1, TOM40, TOM20-2 and RISP (Figure 5.1B). RISP was not efficiently pulled down by MIC60. This may be due to the fact that while the majority of the DGS1 protein co-migrates with complex III (Figure 5.1A), MIC60 was found in a number of protein complexes (Figure 5.1A) (Michaud et al., 2016), and thus only a fraction of the MIC60 antibody recognized MIC60 that was in a complex with RISP. The interaction of MIC60 with the TOM complex is in agreement with a previous report that showed interaction between MIC60 and TOM40 (Michaud et al., 2016). COXII (a subunit of complex V) was not pulled down by either DGS1 antibody or MIC60 antibody, presented as a negative control (Figure 1B). 121 To further confirm the interactions, purified intact mitochondria were treated with membrane-permeable chemical crosslinker disuccinimidyl glutarate to capture transient, semi-stable and stable association of proteins. Disuccinimidyl glutarate is a crosslinker that uses the amine-reactive N-hydroxysuccinimide (NHS) ester group, linking amino to amino groups at < 8-Å in proximity. After crosslinking, centrifugation at 500 RCF for 2 min was performed to pellet and remove aggregated proteins. The supernatant (crosslinked sample) was analyzed by SDS-PAGE and immunodetection. The untreated mitochondria (non-crosslinked sample) were loaded beside as size control. Thus, crosslinked and non- crosslinked samples contained different amounts of mitochondrial input protein and cannot be compared for intensity. Therefore, we rather determined if the target proteins could be cross-linked into a complex with DGS1. DGS1, MIC60, TOM40, TOM20-2 and RISP were crosslinked by disuccinimidyl glutarate in a complex with an apparent molecular mass of 250 kDa (Figure 5.1C, indicated with a red line). TOM40 and TOM20- 2 were also present in another complex with a molecular mass of around 200 kDa, likely representing the TOM complex (Figure 5.1C, indicated with a blue line). Taken together, DGS1 interacts with MIC60, RISP, TOM40 and TOM20-2 forming a multi-subunit protein complex. The dgs1-1 mutation alters the multi-subunit complex To determine the function of the DGS1 protein in the multi-subunit complex, eight different transgenic and mutant lines of Arabidopsis were functionally characterized (Figure 5.2). The dgs1-1 point mutation line (dgs1-1) was from the original study identifying the DGS1 protein (Moellering and Benning, 2010), which has a change in a 122 single amino acid from aspartic acid (D) to asparagine (N) at position 457 close to the predicted transmembrane region (Figure 5.2A). The dgs1 T-DNA insertion line (dgs1-2) containing a T-DNA in the 3rd exon of the dgs1 gene was confirmed by PCR and DNA sequencing (Figure 5.2A) and had a complete loss of DGS1 protein as indicated by immunoblotting (Figure 5.2B). This line was transformed with the sequences encoding the wild type DGS1 and the dgs1-1 mutant protein, respectively, under the control of a 35S CaMV promoter to generate complemented lines with different levels of the native and mutated DGS1 protein. A summary of the mutants/complemented lines is listed in (Supplemental Dataset 2). The DGS1 Comp (L) line produced the DGS1 protein at a low level, half of the DGS1 level in wild-type plants; the DGS1 Comp (H) line produced the DGS1 protein at a high level, more than 10 times of the DGS1 level in wild-type plants (Figure 5.2B); the dgs1-1 Comp (L) expressed the dgs1-1 mutant coding sequence producing the dgs1-1 mutant protein close to the DGS1 level in wild-type plants; dgs1-1 Comp (M) expressed the dgs1-1 mutant coding sequence producing 10 times more dgs1- 1 mutant protein than wild type DGS1 levels; and dgs1-1 Comp (H1) and dgs1-1 Comp (H2) expressed the dgs1-1 mutant coding sequence producing 20 times more dgs1-1 mutant protein than wild type DGS1 levels (Figure 5.2B). To determine whether the abundance of MIC60, TOM40, TOM20s and RISP was affected in these lines, mitochondria were purified from the eight mutant/complementation lines and wild-type plants (Arabidopsis Col-0), and mitochondrial proteins were quantified by immunoblotting. Arabidopsis contains four genes that encode TOM20 proteins, but only three are expressed, TOM20-2, TOM20-3 and TOM20-4 (Lister et al., 2007). The amount of all TOM20 proteins was observed to be reduced by ~ 50% in the dgs1-1 line 123 and dgs1-1 Comp (L) (Figure 5.2B). MIC60 also displayed a 10-15% reduction in abundance in the dgs1-1 line and dgs1-1 Comp (L), but this reduction was only statistically significant in the dgs1-1 Comp (L) line (Figure 5.2B). The other two components of the multi-subunit complex, TOM40 and RISP, remained unchanged in the dgs1-1 line and the dgs1-1 Comp (L) line and were also unchanged in the lines over- expressing wild-type DGS1 protein, indicating that the dgs1-1 mutant protein had a specific effect on MIC60 and TOM20 proteins. Finally, in both the dgs1-1 Comp (M) and (H) lines with high levels of dgs1-1 mutant protein, all the components were reduced in abundance (Figure 5.2B). The abundance of COXII that does not interact with the complex components was not changed in any line (Figure 5.2B). The submitochondrial location of MIC60, TOM40, TOM20-2, RISP and DGS1 was determined by treating intact and outer membrane-ruptured mitochondria with increasing amounts of protease K, with uncoupling protein (UCP) and cytochrome c (Cyt c) as controls for inner-membrane and intermembrane space location, respectively. As expected, UCP was resistant to protease K with intact mitochondria and outer-membrane- ruptured mitochondria, and no difference was observed in the mutant lines when compared to Col-0 (Figure 5.2C). Cyt c was not detected in outer membrane-ruptured mitochondria isolated from Col-0, dgs1-2 and plants complemented with the native DGS1 protein, showing complete rupture of OMM during swelling, with complete release of Cyt c into solution and thus it was absent from the pellet (Figure 5.2C). However, in dgs1-1, dgs1-1 Comp (L), (M) and (H) plants, some Cyt c was still detectable in outer membrane- ruptured mitochondria (Figure 5.2C). This indicates that the OMM is not completely ruptured in dgs1-1, dgs1-1 Comp (L), (M) and (H) lines during the swelling-shrinking 124 procedure. The reason is proposed to be that the structure of MICOS bridging the OMM and IMM has been changed due to the dgs1-1 mutation. Similarly, MIC60 was accessible to protease K in outer membrane-ruptured mitochondria from Col-0, dgs1-2 and plants complemented with the native DGS1 protein (Figure 5.2C), as expected for an intermembrane space exposed protein (Michaud et al., 2016). Similar to Cyt c, MIC60 was still detectable with dgs1-1, dgs1-1 Comp (L), (M) and (H) plants (Figure 5.2C). The opposite was observed with RISP. While it was totally protected from digestion in mitochondria isolated from Col-0, some digestion was observed in the dgs1-1, dgs1-1 Comp (L), (M) and (H) plants upon rupturing the outer membrane (Figure 5.2C). TOM20- 2 and TOM40 are outer-membrane proteins. They were digested by protease K both in intact mitochondria and outer-membrane ruptured mitochondria, and no difference was observed between the lines (Figure 5.S1). DGS1 showed a similar sensitivity as TOM20- 2, with no difference among the lines (Figure 5.2C). The ~ 220 amino acid domain (75- 297aa) at the N-terminal side of the first transmembrane domain, against which the DGS1 antibody was produced, was digested by protease K. This result suggested an N-out and C-out topology for DGS1 with a loop in the intermembrane space and two transmembrane regions (Figure 5.2A and 5.2C). This result also places the mutation in DGS1-1 on the intermembrane space side of the protein, potentially affecting mitochondrial proteins that may interact with DGS1. Thus, the dgs1-1 mutant protein decreases the protease accessibility of MIC60 and Cyt c, while making RISP more accessible, even though the dgs1-1 mutant protein is present at wild-type levels. To determine if the dgs1-1 mutant protein affected the multi-subunit complex containing DGS1, MIC60, TOM40, TOM20s and RISP (Figure 5.1C), mitochondrial 125 proteins from dgs1-1 Comp (L) plants were crosslinked as outlined above. The abundance of DGS1, MIC60, TOM40 and TOM20-2 in the cross-linked 250 kDa complex was significantly reduced in dgs1-1 Comp (L) plants, compared to Col-0 (Figure 5.2D, indicated with red asterisks). In contrast, the abundance of RISP in the cross-linked 250 kDa complex was increased in dgs1-1 Comp (L) plants, compared to Col-0 (Figure 5.2D, indicated with blue asterisks). Based on these results, we conclude that the interaction between DGS1, MIC60, TOM40, TOM20 and RISP proteins is disturbed when dgs1-1 mutant protein is expressed at native levels. The dgs1-1 mutation alters mitochondrial size To determine whether disturbing the multi-subunit complex affects organellar distribution, morphology or size confocal and transmission electron microscopy were carried out. Plant mitochondria exist as a dynamic tubular reticulate network, that is maintained by a balance of fusion and fission (Palmer et al., 2011; Rose and McCurdy 2017). To assess overall morphology in terms of size, constructs for the transient expression of AOX-GFP (targeted to mitochondria) and KDEL-YFP (targeted to the ER) were transiently transformed into protoplasts isolated from Col-0, DGS1 Comp (L), DGS1 Comp (H), dgs1-1 Comp (L), dgs1-1 Comp (M) and dgs1-1 Comp (H1) plants. In Col-0 and plants complemented with the native DGS1 protein, mitochondria surrounded chloroplasts, as is typical in protoplasts (Duchene et al., 2005). In contrast, in dgs1-1 Comp (L) the RFP signal showed that mitochondria were significantly enlarged forming a doughnut shape or oval structures (Figure 5.3A). While these structures varied in size, there were many mitochondria that were much greater in diameter than any mitochondrial 126 structures observed in Col-0, DGS1 Comp (L) and DGS1 Comp (H) lines. With increased levels of the dgs1-1 mutant protein a further increase of mitochondrial size was observed in dgs1-1 Comp (M) and dgs1-1 Comp (H) plants (Figure 5.3A). Enlargement of mitochondria was also observed in yeast mic60 strains and Arabidopsis mic60 knockout lines (Rabl et al., 2009, Michaud et al., 2016). A corresponding reduction in chloroplast size was observed with increasing amounts of the dgs1-1 mutant protein (Figure 5.3A). With dgs1-1 mutant protein levels similar to native levels of the DGS1 wild-type protein, the dgs1-1 Comp (L) line had a significant reduction in chloroplast diameter (Figure 5.3A). By contrast, with higher levels of the dgs1-1 mutant protein, mitochondrial size and diameter were equal to those of chloroplasts in the dgs1-1 Comp (M) and dgs1-1 Comp (H) lines (Figure 5.3A). The YFP signal showed that the ER structure was also affected in dgs1-1 Comp (L), dgs1-1 Comp (M), and dgs1-1 Comp (H) (Figure 5.3A). Together, the shapes of mitochondria, chloroplast and ER were all affected in the lines expressing the dgs1-1 mutant protein. Transmission electron microscopy was carried out to investigate if the internal morphology of mitochondria was altered (Figure 5.3B). As transmission electron microscopy allows morphology to be assessed at a nanometer level it will allow assessment of changes in cristae morphology and/or number. Sections obtained from wild-type plants (Col-0) revealed mitochondria with numerous infoldings of the inner membrane that represent cristae. In contrast, in the dgs1-1 line, dgs1-1 Comp (L) line and dgs1-1 Comp (H1) line, consistently fewer cristae could be detected (Figure 5.3B). The alteration of the inner membrane morphology with fewer cristae provides a possible reason why Cyt c was not completely released from OMM-ruptured mitochondria in the 127 dgs1-1, dgs1-1 Comp (L), (M) and (H) lines observed above (Figure 5.2C). Thus, a change in a MICOS subunit in Arabidopsis results in changes in cristae abundance as described for yeast and mammalian MICOS systems (van der Laan et al., 2016, Schorr and van der Laan, 2018). The dgs1-1 mutation alters mitochondrial lipid composition As MIC60 in Arabidopsis has been shown to play a role in lipid trafficking (Michaud et al., 2016) and the dgs1-1 mutant was identified based on its altered lipid composition (Xu et al., 2008), the lipids of mitochondria and chloroplasts in dgs1-1, dgs1-1 Comp (L), and dgs1-1 Comp (H1) plants were profiled. The chloroplast lipid analysis showed that the total fatty acid profiles of Arabidopsis lines expressing the dgs1-1 mutant protein had a dose- dependent decrease in 16:3 and 18:3 acyl chain abundance and a corresponding relative increase in 16:0, 18:1 and 18:2 acyl chain abundance (Figure 5.4A). The decrease in 16:3 methyl esters abundance can be attributed primarily to the change in monogalactosyldiacylglycerol (MGDG) acyl composition. As the predominant chloroplast lipid, it typically carries 16:3 and 18:3 acyl chains in a 1:2 ratio, respectively, at the sn-1 and sn-2 positions of the glycerol backbone. Additionally, the observed decrease in total abundance of 18:3 acyl chains in the chloroplast lipids was due to a decreased abundance of 18:3 acyl in digalactosyldiacylglycerol (DGDG), phosphatidylglycerol (PG), and sulfoquinovosyldiacylglycerol (SQDG) (Figure 5.4A). In mitochondria, the most abundant lipids, cardiolipin (CL), phosphatidylcholine (PC), and phosphatidylethanolamine (PE), all showed a significant decrease in 18:2 acyl chain abundance that appeared dependent on the abundance of the dgs1-1 mutant 128 protein (Figure 5.4B). Additionally, there was a corresponding increase in the relative amount of 16:0 acyl chains esterified to those lipids (Figure 5.4B). The total lipid profiles showed a significant decrease in the relative amount of CL with increased levels of the dgs1-1 mutant protein, while the relative amount of PC was increased (Figure 5.4B). Analysis of total leaf lipids revealed a dose-dependent decrease in 16:3 acyl chain abundance with a relative increase in 18:1 acyl chain abundance (Figure 5.S2A). The total leaf DGDG and MGDG showed the same trend as in chloroplasts as would be expected. A decrease in the 18:2 acyl chains in PC and 18:3 acyl chains in PG was accompanied by a relative increase in 18:1 acyl chains. For PE a decrease in 18:2 chains was also observed in total leaf lipid profiles like in mitochondria, but an increase in 18:3 acyl chains was observed in total lipids that was not observed for mitochondrial lipids. For PC the same pattern was observed in lipids from mitochondria and total leaf lipids; a decrease in 18:2 acyl chains with a corresponding increase in the 18:1 acyl chains. Thus, the total leaf lipid composition was changed, but the largest effect was observed with mitochondria and chloroplast lipids. It should be noted that in the line with the highest amount of dgs1 mutant protein galactolipids MGDG and DGDG and oligogalactolipids were observed to be associated with the mitochondrial fraction above wild-type background levels (Figure 5.S2B). Typically, MGDG and oligogalactolipids are restricted to chloroplasts. The latter are formed under stress conditions by the action of SFR2 (Moellering et al., 2010). To interpret why the mitochondrial DGS1 protein could affect chloroplast lipid composition, the locations of the DGS1 wild-type protein and the dgs1-1 mutant protein were investigated. The constructs expressing the DGS1 wild-type protein and the dgs1- 129 1 mutant protein tagged with C-terminal GFP or N-terminal GFP were transiently transformed into wild-type protoplasts, with SSU-RFP as chloroplast control, AOX-RFP as mitochondrial control and KDEL-RFP as ER control. The fluorescence imaging revealed that the DGS1 protein was located in mitochondria, while the dgs1-1 mutant protein was associated with both mitochondria and the ER (Figure 5.4C). To confirm these results, chloroplast, mitochondria and ER fractions were isolated from Col-0, dgs- 1, dgs1-2 mutant and complemented lines, followed by immunoblotting with the DGS1 antibody. In Col-0, the DGS1 wild-type protein was only detectable in mitochondria (Figure 5.4D). In contrast, in the mutant lines that expressed the dgs1-1 mutant protein, it was detectable in both mitochondria and the ER (Figure 5.4D), even when expressed at native levels. No DGS1 protein or dgs1-1 mutant protein was detected in chloroplasts (Figure 5.4C and 5.4D). The dgs1-1 mutation affects mitochondrial protein abundance, protein import and alternative respiratory capacity MICOS was reported to affect mitochondrial protein import and respiration in yeast and mammals (Schorr and van der Laan, 2018). To determine whether disturbing the multi-subunit complex by the presence of dgs1-1 mutant protein had the same effects in plants, the abundance of various proteins involved in protein import and respiration was determined. Notably the dgs1-2 null mutant did not affect the abundance of any protein or the activity of the respiratory chain (Figure 5.5A and 5.5B). However, both the dgs1-1 mutant and the dgs1-1 Comp (L) line with protein levels comparable to wild type showed decreased abundance of an outer membrane (cid:0)-barrel protein in plant mitochondria 130 (OM47) (Li et al., 2016), TIM44, and alternative oxidase (AOX) (Figure 5.5A). The consistent decrease in abundance of TOM20s (Figure 5.2B), OM47, TIM44 and AOX (Figure 5.5A) in all lines expressing the dgs1-1 mutant protein indicates that the mutated dgs1-1 protein has an effect on specific proteins or pathways. However, it is worth noting that abundance of almost all mitochondrial proteins tested showed a decrease of various degrees in lines expressing medium and high levels of the dgs1-1 mutant protein (Figure 5.5A). We consider that this is an unspecific effect that may result from the altered mitochondrial lipid content and high level of dgs1-1 mutant protein (Figure 5.4A), and such decreases were not observed in the lines expressing high levels of the DGS1 wild type protein (Figure 5.5A). Along with the alterations in abundance of mitochondrial proteins, a similar trend was observed in respiratory capacity (Figure 5.5B). The activity of the alternative oxidase pathway as assessed by the reduction in n-propyl gallate sensitive oxygen uptake was specifically inhibited by the expression of the dgs1-1 mutant protein (Siedow and Girvin 1980). This reduction in activity of the alternative pathway was observed even when the dgs1-1 mutant protein was expressed at physiological levels. A general reduction in total respiration and respiration through the cytochrome c oxidase pathway, and oxygen consumption driven by oxidation of substrates by complex I and complex II was observed only with high levels of dgs1-1 mutant protein, but not DGS1 wild-type protein (Figure 5.5B). To examine if mitochondrial protein import ability was affected by the presence of the dgs1-1 mutant protein, in vitro import of radiolabeled plant mitochondrial precursor proteins into mitochondria isolated from Col-0, dgs1-1 Comp (L) and dgs1-1 (H1) plants was carried out (Figure 5.5C). For the precursor protein of AOX, representing the general 131 import pathway, and the adenine nucleotide carrier ANT, representing the carrier import pathway, strongly reduced import rates were observed with both dgs1-1 Comp (L) and dgs1-1 Comp (H) plants, compared to Col-0 (Figure 5.5C), indicating an impairment of mitochondrial protein import by the presence of the dgs1-1 mutant protein. Higher drought stress tolerance in dgs1-1 mutant plant lines Quantitative phenotypic analysis revealed that the plants expressing the dgs1-1 mutant protein were smaller than Col-0 and the severity of the retarded growth phenotype was associated with the levels of the dgs1-1 mutant protein (Figure 5.S3). The dgs1-1 Comp (H1) and dgs1-1 Comp (H2) plants, which express high levels of dgs1-1 mutant protein, displayed an early senescence phenotype as evidenced by leaves losing chlorophyll after 3 weeks, a lower number of rosette leaves, and smaller inflourescence (Figure 5.S3). No growth defect was observed for the dgs1-2 null allele, DGS1 Comp (L) and DGS1 Comp (H) plants (Figure 5.S3). Exposing mutant lines to drought stress revealed that dgs1-1, and dgs1-1 Comp (L) were more tolerant than Col-0, dgs1-2, DGS1 Comp (L) and DGS1 Comp (H) plants (Figure 5.6A). The dgs1-1 Comp (H1) plants also displayed higher drought tolerance. To determine the underlying biochemical changes, the rosette leaves #5-7 were subjected to 3,3’-diaminobenzidine tetrahydrochloride (DAB) and nitroblue tetrazolium (NBT) staining to visualize the accumulation of hydrogen peroxide and superoxide radicals, respectively. The dgs1-1, dgs1-1 Comp (L) and dgs1- 1 Comp (H1) lines displayed less ROS accumulation after 8 and 10 days of drought treatment as evidenced by the lower intensity of stains (Figure 5.6A). After 14 days of water deprivation followed by 3 days of recovery after resupply of water, more than 80% 132 of plants survived with dgs1-1, dgs1-1 Comp (L) and dgs1-1 Comp (H) lines, while less than 10% of the Col-0, dgs1-2, DGS1 Comp (L) and DGS1 Comp (H) plants survived (Figure 5.6A). Quantification of relative water content, quantum efficiency of photosystem II (Fv/Fm), total chlorophyll concentration and CO2 fixation (Figure 5.6B) suggested that the plants expressing the dgs1-1 mutant protein were more effective at maintaining water content and photosynthetic efficiency during drought stress. DAB staining of the dgs1-1 lines revealed that the dgs1-1 Comp (H1) plants, which displayed an early senescence phenotype (Figure 5.S3), contained higher levels of H2O2 than Col-0 plants even when not subjected to adverse (drought) conditions (Figure 5.6A, bottom panel), yet these plants displayed survival under drought compared to lines expressing the DGS1 native protein. Quantification of H2O2 in leaves revealed that plants with high levels of the dgs1-1 mutant protein contained higher levels of endogenous H2O2 (Figure 5.S4). This is consistent with the original identification of dgs1-1 where over- expression resulted in growth defects and an increase of two to three-fold in H2O2 (Xu et al., 2008). The dgs1-1 and dgs1-1 Comp (L) plants, expressing lower level of dgs1-1 mutant protein, contained a similar level of H2O2 as Col-0 plants and did not show acceleration of dark-induced leaf senescence (Figure 5.S4). Thus, the amount of the dgs1-1 mutant protein present is important in determining the phenotypes observed. To determine the underlying molecular alterations in dgs1-1 and dgs1-1 Comp lines, transcriptomes of mutant and wild-type plants were analyzed by RNA-Sequencing. The transcriptome of plants grown under non-limiting growth conditions was not severely altered in the mutant lines, except for the complemented lines with high levels of DGS1 wild-type protein or dgs1-1 mutant protein (Figure 5.7A, 5.7B). DGS1 Comp (H) and dgs1- 133 1 Comp (H1) had 52 and 162 genes with altered basal expression, respectively, while all other lines had less than 10 differentially expressed genes (DEGs) (fold change > 2 and FDR < 0.05) compared to the wild type (Supplemental Dataset 3). Under water limited conditions, the transcriptome of plants containing the dgs1-1 mutant protein different to wild type and dgs1-2 lines complemented with the wild type coding sequence for DGS1. Compared to the wild-type that had 2230 and 2848 genes up- and down-regulated by drought, plants showed a reduced number of DEGs, with 1884, 1740 and 1640 genes up- regulated and 2450, 2333 and 2222 genes down-regulated in dgs1-1, dgs1-1 Comp (L) and dgs1-1 Comp (H1), respectively (Figure 5.7A). From the total number of drought- responsive genes in the wild type, only 65%, 66% and 55% were similarly (> 2-fold change in the same direction and FDR < 0.05) responsive in dgs1-1, dgs1-1 Comp (L) and dgs1-1 Comp (H1), respectively. To further reveal the effect of the dgs1-1 allele on the drought-responsive processes, genes that were commonly altered in dgs1-1 and dgs1-1 Comp (L) were analyzed as both these lines provide a read-out of the effects of the dgs1-1 allele expressed at native levels (Figure 5.7B, Supplemental Dataset 3). From the 720 DEGs in dgs1-1 and dgs1-1 Comp (L) compared to Col-0 under drought stress, three clusters could be identified that are indicated in blue, red and green (Figure 5.7B). Genes in each cluster were functionally classified by GO term enrichment analysis as indicated with the same colors (Figure 5.7C). Two hundred and sixty-five genes (blue cluster) were drought induced in Col-0, dgs1-2, DGS1 Comp (L) and DGS1 Comp (H), but their induction was partially or completely abolished in dgs1-1 mutant lines (Figure 5.7B). These genes are associated with GO terms related to abiotic stresses such as drought, salt, hydrogen 134 peroxide, cold and stress-induced senescence, indicating that all the DGS1-1 expressing plants have largely not activated abiotic stress responses (Figure 5.7C). Among them are genes encoding transcription factors for drought-induced gene expression (ANAC019, ANAC072 and ANAC032), proteins involved in osmoprotection (EARLY RESPONSE TO DEHYDRATION SIX-LIKE 1, ARABIDOPSIS MITOCHONDRIAL BASIC AMINO ACID CARRIER 2, and LATE EMBRYOGENESIS ABUNDANT and DEHYDRIN family proteins) and proteins involved in cell wall remodeling (GLYCOLIPID TRANSFER PROTEIN, XYLOGLUCAN ENDOTRANSGLUCOSYLASE/HYDROLASE, and CELLULOSE SYNTHASE family proteins). A second cluster (red) consists of 367 genes involved in photosynthesis, primary metabolism and auxin signaling. These are down- regulated in Col-0, dgs1-2, DGS1 Comp (L) and DGS1 Comp (H) plants during drought stress conditions. This re-direction of energy/growth metabolism during drought stress was severely diminished in all lines expressing the mutated dgs1-1 protein, again consistent with a lack of stress response. A third cluster (green) contains genes that are up-regulated in the three dgs1-1-expressing genotypes only. These are involved in sulfur metabolism and glucosinolate biosynthesis pathways (METHYLTHIOALKYLMALATE SYNTHASE1, METHIONINE AMINOTRANSFERASE4, CYTOCHROME P450 79F1, 3- ISOPROPYLMALATE DEHYDRATASE SMALL SUBUNIT 1 and the MYB28, MYB29 and MYB76 transcription factors). Together, these data indicate that the dampened transcriptomic responses to drought stress in all genotypes expressing dgs1-1 are attributed to the increased drought tolerance, which are probably caused by the altered physiological characteristics (e.g. lipid composition) or the ectopically expressed genes, e.g. involved in glucosinolate biosynthesis. 135 Phylogenetic and co-expression networks of DGS1 putative orthologs As DGS1 was putatively orthologous to NCA2 of yeast, a comparative analysis of their phylogeny was performed. DGS1/NCA2 orthologues were detected in plants and fungi and in a limited number of animal groups, in Amoebozoa, Chromists and Excavates, but not the Metazoa. It was also absent from the Alveaeolata (Figure 5.8A, Supplemental Dataset 4). However, the widespread presence in plants, fungi and the Choanoflagellata (Salpingoeca rosetta), Filasterea (Capsaspora owcazarzaki) and Ichthyosporea (Sphaeroforma arctica) suggests that it was lost in multi-cellular animals. To further explore their relationship, DGS1 and NCA2 were analyzed for co- expressed genes (Figure 5.8B, Supplemental Dataset 5). DGS1 was strongly co- expressed with MIC60 (15th most co-expressed genome wide), but not with MIC10, whereas NCA2 was co-expressed with MIC27, MIC26, MIC60, and MIC12. Both DGS1 and NCA2 were co-expressed with genes related to mitochondrial fission and morphology. These included the Arabidopsis gene encoding dynamin-related GTPase DRP3B and its yeast ortholog Dnm1. Those together with yeast mitochondrial fission 1 (Fis1) are required for mitochondrial and peroxisome fission (Otsuga et al., 1998; Mozdy et al., 2000; Bleazard et al., 1999; Fujimoto et al., 2009). Arabidopsis drp3b mutants and yeast dnm1 and fis1 mutants are defective in mitochondrial fission and display altered mitochondrial morphology, similar to the dgs1-1 mutant. Interestingly, cardiolipins are required for mitochondrial fission by stabilizing the Arabidopsis DRP3 oligomer complexes (Pan et al., 2014). Similarly, a functional MICOS and cardiolipins are required for Drp1 complex stability and mitochondrial morphology in mammalian cells (Bustillo- 136 Zabalbeitia et al., 2014, Pan et al., 2014). In addition, DGS1 was co-expressed with MIRO-RELATED GTP-ASE 1 that is involved in mitochondrial motility and morphology, in a pathway independent of the DRP3 fission machinery (Yamaoka and Leaver, 2008). NCA2 was co-expressed with various genes involved in lipid metabolism and transport. For instance, the OMM protein Mitochondrial distribution and morphology 34 (Mdm34) is a core component of the ER-mitochondrial tethering (ERMES) complex and functions in mediating lipid import from the ER into the mitochondria (Kornmann et al., 2009, Klecker et al., 2015). Translocator, assembly and maintenance 41 (TAM41), a mitochondrial phosphatidate cytidylyltransferase is required for cardiolipin biosynthesis (Kutik et al., 2008), and Cardiolipin-specific deacylase 1 (CLD1) is a mitochondrial cardiolipin-specific phospholipase (Beranek et al., 2009). DGS1 was co-expressed with PHOSPHOLIPASE D P1 that is required for extraplastidial galactolipid biosynthesis during phosphate starvation (Cruz-Ramirez et al., 2006). In addition, NCA2 (but not DGS1) was co- expressed with genes encoding several components of the mitochondrial complex III, complex IV, and the F1F0-ATP synthase complex. These results showed that DGS1 and NCA2 were co-expressed with conserved cellular processes, indicating a similar function across phyla. Discussion A number of lines of evidence indicate that the outer mitochondrial membrane protein DGS1 from Arabidopsis forms a multi-subunit complex that contains MIC60, a conserved core subunit of MICOS and previously identified in Arabidopsis as an important component involved in lipid trafficking in a complex called MTL (Michaud et al., 2016) 137 (Figure 5.8C). Consistent with our results, TOM40 and TOM20 proteins and RISP were also identified as components of MTL previously (Michaud et al., 2016), and our work establishes a link between DGS1 and this complex. We found that DGS1 represented an outer membrane component of MICOS that played a role in lipid homeostasis in plant organelles, possibly by forming contact sites with the ER and/or chloroplast/plastids to facilitate lipid import into plant mitochondria. A complete loss of the DGS1 protein in dgs1- 2 had no apparent effect on mitochondrial activity, mitochondrial protein composition and size. By contrast, the presence of the dgs1-1 mutant protein resulted in a number of alterations. At a comparable molecular level, the introduction of the dgs1-1 mutant protein resulted in decreased amounts of TOM40, TOM20 and MIC60 in this MICOS complex (Figure 5.2D) and weakened protease K sensitivity of MIC60 in our experiments (Figure 5.2C), while the increased amounts of the RISP (Figure 5.2D) and its increased sensitivity to protease K (Figure 5.2C), indicate an altered structure of the MICOS complex (Figure 5.8C). This is likely the consequence of the point mutation present at the intermembrane space side of the dgs1-1 protein, altering its binding affinity with these proteins (Figure 5.8C). Notably, a decrease in the amount of TIM44 in mitochondria was evident (Figure 5.5A), likely being responsible for the reduced rates of protein import observed (Figure 5.5C). In plants, the general mitochondrial processing peptidase (MPP) is integrated into the cytochrome bc1 complex (Braun et al., 1992, Glaser et al., 1994), and therefore an increased association of the dgs1-1 mutant protein with the RISP may destabilize TIM44. Alternatively, altered mitochondrial membrane protein abundances could be due to the altered lipid composition of mitochondria, with fewer cristae present in mitochondria that contain the dgs1-1 mutant protein (Figure 5.3B). The non-bilayer-forming 138 phospholipids CL and PE are required for cristae structures and stabilization of respiratory complexes, whereas bilayer-forming phospholipid PC stabilizes TIM complexes (Schuler et al., 2016). Together, alterations in lipid composition, protein abundance and potentially also altered mitochondria-ER contacts (that are hotspots for mitochondrial fission) by dysfunctional MICOS may be part of the reasons for the altered organelle size of mitochondria, chloroplasts and the ER as evidenced by visualization with fluorescence tagging of proteins (Figure 5.3A). The significantly altered size of mitochondria that express the dgs1-1 mutant protein at physiological levels suggests that the balance between fusion and fission is altered in the dgs1-1 mutant lines. This may result from an interaction of DGS1 with proteins involved in mitochondrial fusion and fission and mitochondrial dynamics (Figure 5.8B). Alternatively, cardiolipins which were altered in dgs1-1 mutant lines stabilize Arabidopsis DRP3 that is required for fission (Pan et al., 2014). While the majority of DGS1 co-migrated with MIC60 in complex III on BN-PAGE, it was clear that MIC60 was present in a number of other complexes including supercomplexes I+III, complex I, III and V (Figure 5.1A). Previously, it was reported that the MTL complex in Arabidopsis is comprised of 200 proteins, including DGS1 (Michaud et al., 2016). Thus, it is likely that the interaction of DGS1 with MIC60 is a dynamic interaction, positioning MIC60 at the interface with protein and lipid import into plant mitochondria. Notably the impact of the dgs1-1 mutant protein on protein and lipid import into mitochondria, directly or indirectly decreasing the abundance of both TOM20s and TIM44, indicates that mitochondrial biogenesis is affected by disordering the stability or dynamics of this complex. While MIC60 and MIC10 seem to be the only subunits 139 conserved between yeast, plants and animals, alteration of the MICOS complex appears to have very similar impacts in all three phyla. MICOS in yeast and mammals has multi- faceted effects on mitochondrial morphology, protein import and abundance, oxidative phosphorylation and lipid synthesis (van der Laan et al., 2016, Schorr and van der Laan, 2018), which are similar to the alterations observed in this study with the expression of the dgs1-1 mutant protein. DGS1 in plants displays sequence similarity with a yeast protein that has been called Nuclear Control of ATP Synthase 2 (NCA2), identified in a genetic screen for genes required for the assembly/accumulation of the ATP synthase complex in yeast by yet unknown mechanisms (Pelissier et al., 1995). A comprehensive analysis of the yeast mitochondrial proteome identified it as an outer membrane protein (Zahedi et al., 2006). DGS1/NCA2 proteins are present in a wide range of eukaryotes, but appear to have been lost in Metazoa (Figure 5.8A). The DGS1/NCA2-encoding genes also displayed similarities in their co-expressed genes that encode MICOS proteins, but also proteins involved in mitochondrial fission, morphology and lipid homeostasis (Figure 5.8B). These proteins are conserved across kingdoms and act at mitochondria-ER contact sites where mitochondrial fission is initiated (Friedman et al., 2011). We propose that the DGS1 and NCA2 proteins play similar, but not identical, roles in yeast and plants for lipid import. This hypothesis is based on the fact that the decrease in ATP synthase in yeast is likely a secondary effect of an altered lipid composition, and cardiolipin may be a specific target that is required for the stability of the ATP synthase complex (Mehdipour and Hummer, 2016). To be noted, DGS1 and NCA2 also have specific functions. Plants carrying a homozygous null allele dgs1-2 had no detectable alterations in plant growth or 140 mitochondrial function. Thus, while a specific point mutation of DGS1 in dgs1-1 resulted in an altered phenotype, a complete knockout can be compensated by other routes. This suggests that while DGS1 is part of the plant MICOS complex, import of lipids into mitochondria can still take place in the absence of the DGS1 protein. It should also be noted that it is possible that DGS1 in plants has developed additional functions needed for the complex interaction of plants with the changing environment. AOX is a well-established marker of mitochondrial retrograde signaling (Ng et al., 2014) as it is induced upon perturbation of mitochondrial functions. An original observation for the dgs1-1 line was that the amount of AOX in mitochondria was reduced (Moellering and Benning, 2010). Altered AOX levels are likely attributed to altered lipid membrane homeostasis and/or membrane morphology affecting protein stability. However, as AOX is typically induced by a variety of abiotic stresses (Ng et al., 2014), and the drought stress responsive transcriptome was affected by the dgs1-1 allele, altered signaling in dgs1-1 mutant cannot be excluded. Although most of the altered responses represent a dampened stress response due to the increased drought tolerance of dgs1-1 plants, a small set of genes involved in sulfur metabolism and glucosinolate biosynthesis was mis-expressed in these lines (Figure 5.7B). Up-regulation of glucosinolate biosynthesis and metabolism has been linked to responses to drought stress (Eom et al., 2018). A positive regulator of glucosinolate metabolism in Arabidopsis, MYB29, has been previously shown to be a negative regulator of AOX expression (Zhang et al., 2017), and this may account for the reduction of AOX observed in the dgs1-1 lines (Figure 5.5A). This supports the suggestion that there is an antagonistic relationship in the induction of glucosinolate metabolism and AOX (Zhang et al., 2017), with the former 141 favored in the dgs1-1 lines. Glucosinolate biosynthesis is also regulated by jasmonic acid and there is an antagonistic interaction between the jasmonic and salicylic acid hormone signaling pathways in Arabidopsis (Thaler et al., 2012). Salicylic acid is involved in the induction of AOX, and research has indicated that this acts at the post-transcriptional level (Rhoads and McIntosh, 1992, Ho et al., 2008). Thus, the reduction in AOX may be due to altered plant hormone signaling. Additionally, the GENOMES UNCOUPLED 4 (GUN4) gene was greatly up-regulated upon drought in the dgs1-1 background (Figure 5.7B). GUN4 plays a regulatory role in promoting chlorophyll accumulation in response to changing environmental conditions (Larkin et al., 2003). This is consistent with the observed drought-resistant phenotype, with chlorophyll concentrations and photosynthetic capacity maintained longer in dgs1-1 mutant lines than in the wild type. Materials and Methods Plant Materials The Arabidopsis DGS1 (AT5G12290) mutant dgs1-2 (SAIL_391_F04) and dgs1-1 alleles were described previously (Xu et al., 2008, Moellering and Benning, 2010). All complementation lines, DGS1 Comp (Low/High) with low (L) or high (H) expression of native DGS1 protein and dgs1-1 Comp (Low/ Middle /High) with low (L), moderate (M) or high (H) expression of the dgs1-1 mutant protein were generated in the null allele dgs1- 2 background by gateway cloning of the DGS1 or dgs1-1 coding sequence into the binary destination vector pK7WG2 and Agrobacterium-mediated transformation via floral dipping. The cloning primers are listed in (Supplemental Dataset 6). dgs1-1 Comp (H1) 142 and dgs1-1 Comp (H2) are two individual complementation lines with high but nearly equal levels of dgs1-1 mutant protein. The mutants and complementation lines were confirmed by quantification of the DGS1 or dgs1-1 mutant protein. Plant Growth and Treatments For normal conditions, Arabidopsis plants were grown in growth chambers at 22°C, 100 µmol m-2 s-1 light (color code 840, light color 4000K – cool white, 5240 Lumen and 150 cm long, Philips Master TL-D 56 W / 840 RefleX) in a 16-h light/8-h dark photoperiod. Seeds were sterilized and sown on Gamborg’s B5 medium (PhytoTechnology, Austratec) supplemented with 3% (m/v) sucrose, 0.43 g/L Gamborg’s B5 salts (Austratec), 2 mM MES hydrate (Sigma-Aldrich), and 0.90% (w/v) Difco™agar (BD Biosciences). The pH was adjusted to 5.7. Seeds were stratified for 48 h before being transferred to growth chambers in all experiments. For soil-based phenotyping or drought treatment, all lines were grown in a randomized design on soil mix. Watering was withheld when plants were 24 days old and plants were resupplied with water after 14 days. Isolation of mitochondria, chloroplast and ER Mitochondria, chloroplasts and ER were isolated as described (Cline et al., 1981, Bessoule et al., 1995, Lister et al., 2007), respectively, from seedlings grown on Gamborg’s B5 medium for 2 weeks. Fractions were stored at -80°C and maintained on ice when in use. 143 Blue-native PAGE Twenty micrograms of mitochondria was solubilized in 2 µl 5% (w/v) digitonin and separated on a NativePAGE™ Novex® 4-16% Bis-Tris Gel (Life Technology, Melbourne, Australia). One-dimensional BN-PAGE was carried out as described previously (Eubel et al., 2005). Following separation, gels were fixed and stained with Coomassie colloidal dye (Bio-Rad) or transferred to a PVP membrane followed by immunodetection with the antibodies of DGS1, MIC60, TOM40, RISP and COXII detailed in Supplemental Dataset 7. Immunoprecipitation (IP) Two hundred micrograms of mitochondrial proteins was solubilized and immunoprecipitated using the Immunoprecipitation Kit (Protein A) (Roche, Germany). Sample buffer was added after wash and denaturing of proteins was achieved by heating to 100°C for 3 min that was followed by 30 seconds centrifugation at 12,000 RCF at 25°C to release proteins from beads. Samples were resolved by SDS-PAGE, transferred to a nitrocellulose membrane, and immunodetected with the antibodies of DGS1 and MIC60 detailed in Supplemental Dataset 7. Cross-Link Assay Protein cross-linking using disuccinimidyl glutarate was performed according to manufacturer’s instruction (Thermo Scientific, Melbourne, Australia). One milligram of freshly isolated mitochondria from 2-week-old water cultured Arabidopsis wild-type and mutant plants were precipitated by centrifugation at 17,500 RCF at 4°C for 2 min and 144 resuspended with PBS buffer (100 mM sodium phosphate, 0.15 M NaCl, pH 7.2) to a final concentration of 2 µg/ µl. The samples were subsequently incubated with 5 mM disuccinimidyl glutarate (DSG)-PBS buffer. Samples were placed on ice for 2 h and quenched with 1 M Tris-HCl (pH 7.5) for 15 min at room temperature. Centrifugation at 500 RCF at 4°C for 2 min was performed to pellet and remove aggregated proteins. The supernatant was kept for further analysis by SDS-PAGE and immunodetection. Generation of the DGS1 antibody The fragment of the DGS1 protein (amino acids 75–297) before the first predicted transmembrane-spanning domain (TM) was cloned by PCR reactions using primers listed in Supplemental Dataset 6, cloned into the Gateway pDEST17 vector (Invitrogen, Sydney, Australia), and transformed into the E.coil BL21-AI™ (DE3) expression strain according to the manufacturer’s instructions (Invitrogen, Sydney, Australia). A single colony was cultured in 50 ml of LB medium containing 50 µg ml-1 carbenicillin, 0.1% (m/v) glucose and 0.05% (m/v) L-arabinose with shaking overnight at 37°C until the OD600 reached 0.6–1.0. Afterwards, 950 ml of fresh LB medium was inoculated and cells were grown with shaking until the OD600 reached 0.4. Protein expression was induced with 1.0 mM isopropyl-β-D-thiogalactopyranoside (IPTG) and the cell culture was harvested after 3 h incubation by centrifugation at 8,000 RCF at 4°C for 20 min. Cells were resuspended in 50 ml wash buffer (50 mM KH2PO4, 300 mM KCl, 5 mM imidazole, pH 8.0) and lysed by sonication. The protein was purified by denaturing immobilized metal affinity chromatography (IMAC) using the Profinia protein purification system (Bio-Rad, Sydney, 145 Australia). Three aliquots of 200 µg purified protein were used for injection into rabbits (WEHI, Australia) and the serum was tested by immunoblot. Immunoblot Analysis Proteins of different cell fractions were resolved by SDS-PAGE and transferred to a Hybond-C extra nitrocellulose membrane. Immunodetections were performed as described previously (Wang et al., 2012). The antibodies used in this study were listed in Supplemental Dataset 7. The intensities of bands of interest were quantified using Image Lab™ software (Bio-Rad) and calculated relative to the wild type. Three biological replicates were performed. Significant difference was determined using a Student’s t-test and P ≤0.05 is indicated by asterisks or numbers in red. Antibodies used in this study are listed in Dataset S7. Outer membrane ruptured mitochondria preparation and protease treatment Two hundred micrograms of freshly isolated mitochondria were precipitated, and the pellet was gently resuspended in 20 μl SHE buffer (250 mM sucrose, 1 mM EDTA and 10 mM HEPES-KOH, pH 7.4) and incubated on ice for 15 min after mixing with 310 μl 20 mM HEPES-KOH (pH 7.4). Following incubation, 50 μl 2 M sucrose and 20 μl 3 M KCl were added to rupture the outer membrane. Equal amounts of outer membrane ruptured mitochondria and intact mitochondria were incubated for 30 min on ice with 0 to 2 mg/ml protease K, and 4 μl of 100 mM phenylmethane sulfonyl fluoride (PMSF) was added to terminate the reaction. Samples were precipitated by centrifugation at 13,000 RCF at 4°C for 3 min and, resuspended in 100 µl of SDS-PAGE sample buffer (10% (m/v) SDS, 1% 146 (v/v) β-mercaptoethanol, 18.75% (v/v) glycerol, 0.1% (m/v) bromophenol blue and 150 mM Tris-HCl, pH 6.8). Ten microliters of each sample was used for SDS-PAGE and immunodetection. Protoplast Preparation and Fluorescence Microscopy Protoplasts were isolated from the true leaves of 4-week-old Arabidopsis plants using Tape-Arabidopsis Sandwich methods as described (Wu et al., 2009). A mixture of protoplasts (2-5 × 105 cells/ml) and 20 μg of plasmid DNA was incubated with 0.2 ml 40% (m/v) PEG 4000 at room temperature for 20 min, precipitated at 100 RCF for 1 min and resuspended with 1 ml W5 solution. Washes were repeated twice and protoplasts incubated in the dark at 19-20°C overnight. Fluorescence imaging was carried out using a Zeiss LSM780 laser scanning confocal microscope with an LD C-Apochromat 40x/1.1 water or 100x/1.4 Oil-immersion objective in multi-track channel mode. Image processing was performed using ZEN 2.3 (blue edition, Carl Zeiss Microscopy GmbH, 2011). Electron Microscopy Leaf tissue samples were cryofixed using a Leica EM ICE high pressure freezer (Leica Microsystems) followed by freeze substitution in 1% osmium tetroxide in acetone for 72 h at -85oC. The samples were then slowly warmed to room temperature over the course of 24 h. The fixative was removed and samples were washed three times with acetone before infiltration and embedding with Spurr’s epoxy resin. Thin (70 nm) sections were cut using a Leica UC7 microtome and the samples were imaged using a Biotwin CM120 or FEI Spirit transmission electron microscope (Thermo Fisher Scientific). 147 Lipid Extractions and Analysis Lipids were extracted from isolated Arabidopsis mitochondria and chloroplasts, and prepared for gas-liquid chromatography (GLC) following (Wang and Benning, 2011). Mitochondrial phospholipids were separated using a silica TLC plate treated with 2.3% boric acid and a chloroform:ethanol:water:triethylamine solvent system (30:35:7:35, v/v). Lipid bands were visualized with a non-destructive primuline stain. Chloroplast polar lipids were separated using a silica TLC plate (Si250 with pre-adsorbent layer, Mallinckrodt Baker, NJ) treated with (NH4)2SO4 and an acetone: toluene: water (91:30:7, v/v) solvent system. Lipid bands were visualized with brief iodine vapor staining. Individual lipids were scraped, and their fatty methyl ester profiles analyzed using GLC (Xu et al., 2003). Measurements of Respiratory Parameters on Plant Mitochondria Oxygen uptake by isolated mitochondria was measured as described previously (Jacoby et al., 2015), using a Clark-type O2 electrode (Hansatech, United Kingdom). Oxygen consumption driven by NADH oxidation was measured by adding malate (5 mM) and NADH (1 mM). Malate generates NADH in the mitochondrial matrix via oxidation of malate in the tricarboxylic acid cycle and added NADH can be oxidized by the alternative external NADH dehydrogenases (Palmer and Passam1971; Coleman and Palmer 1972; Palmer and Ward 1985). Rotenone (5 μM) was added to specifically inhibit complex I to determine complex I NADH driven oxygen consumption, and the activity remaining after rotenone addition represents oxygen consumption driven by the internal and external NADH dehydrogenases. Complex II-driven oxygen consumption was assessed by adding succinate (5 mM) and assessing the malonate (5 mM) sensitive oxygen consumption rate, 148 with malonate acting as a complex II inhibitor. Activity of the cytochrome c oxidase pathway was measured in a 1 ml reaction volume of aerated respiration medium containing 0.3 M sucrose, 5 mM KH2PO4, 10 mM TES, 10 mM NaCl, 2 mM MgSO4 and 0.1% (w/v) BSA (pH 7.2), in the presence of saturating concentrations of succinate (5 mM) and NADH (1 mM), ATP (0.5 mM), ADP (0.3 mM), malate (10 mM), pyruvate (10 mM), coenzyme A (12 μM), thiamine pyrophosphate (0.2 mM) and NAD+ (2 mM). Potassium cyanide (KCN, 1 mM) and n-propylgallate (nPG, 0.5 mM) were used to inhibit complex IV and AOX, respectively. Data were analyzed based on three biological replicates. Statistical evaluations were conducted by means of the two-way ANOVA with post-hoc Tukey HSD test integrated in GraphPad Prism 7 (GraphPad Software Inc.). Differences with P-value < 0.05, P-value < 0.01 and P-value < 0.001 were considered as significant and indicated as *, **, and ***, respectively. In Vitro Mitochondrial Protein Import [35S]-Met–labelled precursor proteins were synthesized and imported into freshly isolated mitochondria as described previously (Lister et al., 2007). Equal quantities of mitochondria from different genotypes were used for import reactions. Following import, mitochondria were precipitated at 20,000 RCF for 5 min and subjected to SDS-PAGE. Gels were stained in Coomassie Brilliant Blue, dried, and exposed to a BAS TR2040 phosphor-imaging plate (Fuji) for 24 h. The exposed plate was visualized using the BAS 2500 Bio-Imaging Analyzer (Fuji). 149 In Situ Detection of Reactive Oxidative Species (ROS) Detection of H2O2 by 3,3-diaminobenzidine staining was carried out as previously described (Förster et al., 2005) and reactive oxygen species in the form of O2•− were determined by staining with nitroblue tetrazolium as described (Sedigheh et al., 2011). Detached leaves were incubated in freshly prepared 1 mg/ml DAB solution (pH 7.0) or 600 μM NBT solution (pH 7.0) at room temperature in the dark (DAB stain 8 h, NBT stain 4 h), and then transferred to destaining solution (ethanol: acetic acid = 3:1, v/v) to remove the chlorophyll. The bleaching solution was changed every 12 h until all chlorophyII was removed. Leaves were fixed in 50% (v/v) glycerol and scanned. Three leaves (true leaves 5, 6, and 7) per plant and three plants per genotype were assayed for both H2O2 and O2•− detection. Dark-induced Senescence Arabidopsis plants were grown on soil in growth chambers at 22°C, 100 µmol m-2 s-1 light intensity in a 16-h light/8-h dark photoperiod. For dark-induced senescence, the true leaves 6 and 7 from 3-week-old Arabidopsis plants were covered with foil for 5 days. The control plants were not covered with foil, and were kept growing in a 16-h light/8-h dark photoperiod. Quantification of H2O2 H2O2 was quantified as described previously (Liu et al., 2010), using leaf extracts from 3- week-old plants. The extract was diluted accordingly and then used for H2O2 determination with an Amplex Red Hydrogen Peroxide/Peroxidase Assay Kit 150 (Thermofisher, Melbourne, Australia). The values were obtained from three biological replicates, and each replicate was a pooled sample of the sixth and seventh leaves from one plant. The data were analyzed by analysis of variance (ANOVA), and means were compared using a Student’s t-test. Relative Water Content Analysis The relative water content of each genotype was measured as described previously (Giraud et al., 2008). Five biological replicates were measured for each genotype and each replicate contained the whole rosette leaves from one plant. Student’s t-test was performed to determine significant differences (P ≤ 0.05). Chlorophyll analysis and fluorescence The total chlorophyll content was measured as described previously (Li et al., 2016). The chlorophyll fluorescence Fv/Fm (maximum quantum yield of PSII) was pulsed with 120 μmol m−2 s−1 of actinic light using the IMAGING-PAM M-series Chlorophyll Fluorescence System (Walz) after a 20 min dark acclimation as previously described (Rossel et al., 2006). At least three biological replicates were measured for each genotype and each replicate was a pooled sample of true leaves 5, 6 and 7 from one plant. Student’s t-test was performed to determine significant differences (P ≤ 0.05). 151 Gas Exchange Individual leaves were enclosed in a 1 cm reach chamber (Li-COR, 6400-15) that was attached to a Li-COR portable photosynthesis system (BioScientfic Ltd) according to (Li et al., 2014) with some modifications. Net photosynthetic CO2 fixation rate was measured at 22°C, 400 μmol m−2 s−1 PPFD RGB light source (Li-COR, 6400-18A), 400 μmol mol−1 CO2 concentration, and 65–75% relative humidity. The rate of respiration was determined by measuring the net photosynthetic CO2 fixation rate in darkness. The true leaves 5, 6, and 7 from each plant, and 5 plants from each genotype were measured. Student’s t-test was performed to determine significant differences (P ≤ 0.05). Total RNA Preparation and cDNA Synthesis Total RNA was isolated in biological triplicates from pooled #5, 6 and 7 true leaves from 3-week-old plants using the Sigma Spectrum™plant total RNA isolation kit (Sigma- Aldrich) according to manufacturer’s instructions. On column DNase treatment was carried out to remove genomic DNA (Thermo Scientific™). The cDNA was synthesized using the iScript™ cDNA Synthesis Kit (Bio-Rad, Hercules, CA, USA) according to the manufacturer’s instructions. Global Transcript Analysis Total RNA was isolated in biological triplicates from pooled #5, 6 and 7 true leaves of well-watered and 8-day-drought-treated wild-type (Col-0) and mutant plants after growth under normal conditions for 3 weeks. RNA isolation was performed using the RNeasy Plant mini kit and DNA was removed via on-column DNase digestion using the RNase- 152 Free DNase kit according to the manufacturer’s instructions (Qiagen, Sydney, Australia). The RNA was eluted in molecular grade DNase- and RNase-free water and the integrity was validated using a TapeStation 2200 system (Agilent, Mulgrave, Australia). RNA-seq libraries were prepared using the TruSeq Stranded mRNA Library Prep Kit according to the manufacturer’s instructions (Illumina) and sequenced on a HiSeq1500 system (Illumina) as 60 bp reads or a NextSeq550 system as 75 bp reads with an average quality score (Q30) of above 95%. The raw reads (on average 19.7M per sample) were quality controlled by FASTQC (v 0.11.5, https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and trimmed by trimgalore (v 0.4.5, https://github.com/FelixKrueger/TrimGalore). Clean reads were pseudo-aligned to the representative transcript models of the Arabidopsis Araport 11 annotation (Cheng et al., 2017) using Kallisto (v 0.43.1) to generate TPM/counts tables (Bray et al., 2016). Next, glmFit function in edgeR (Robinson et al., 2010) was used to fit the negative binomial generalized linear model (GLM) and glmLRT was used to carry out the likelihood ratio test. Differentially expressed genes were identified with a threshold of fold change > 2 and FDR < 0.05. For the Gene Ontology enrichment analyses, the Cytoscape (v3.5.1) (Shannon et al., 2003) plug-in ClueGO (Bindea et al., 2009) was used. CluoGO was run with default settings, except for setting the P value cutoff < 0.1, GO tree interval level to 4–9, and ≥ 3 or more genes from either cluster associated to a GO term, representing ≥ 4% (for clusters Blue and Red) or ≥ 2% (for cluster Green) of the associated GO term genes. All sequencing data have been submitted to the NCBI SRA archive under Project ID PRJNA475427. 153 Mass Spectrometry Preparation of samples from BN gel bands Individual samples were excised from BN gels and destained (50 mM ammonium bicarbonate/acetonitrile) overnight. Proteins were reduced in 1 µl of 200 mM tris(2- carboxyethyl)phosphine (TCEP) in 100 μl of water for 1 h. After washing the gel bands (50 mM ammonium bicarbonate/acetonitrile), samples were alkylated with 4 µl of 1 M iodoacetamide (IAA) in 100 µl of water for 30 min in the dark. Proteins were then digested with trypsin (0.2 μg trypsin (Promega Sequencing Grade) for 16 h at 37°C) and peptides collected after extraction (70% acetonitrile/water). LC-MS/MS analysis Peptides were reconstituted in 0.1% TFA and 2% acetonitrile (ACN) and loaded onto a C18 PepMap 100 µm ID × 2 cm trapping column (Thermo-Fisher Scientific) at 5 µl/min for 6 min, and washed for 6 min before switching the pre-column in line with the analytical column (PepMap RSLC C18, 2 μm, 75 μm i.d. x 25 cm, Thermo Fisher Scientific). Peptides were loaded and separated for 60 min. The separation was performed at 250 nl/min using a non-linear ACN gradient of buffer A (0.1% formic acid, 2% ACN) and buffer B (0.1% formic acid, 80% ACN), starting at 5% buffer B to 55% over 55 min, then 100% buffer B for 5 min followed by an equilibration step of 15 min (0.1% formic acid, 2% ACN). Data were acquired using the Xcalibur software v4.1 (Thermo Fisher Scientific). The mass spectrometer was programmed to acquire in a data-dependent mode using a maximum ion injection time of 50 ms A. Full scans were acquired in the Orbitrap mass analyzer with a resolution of 120,000 at 200 m/z (3E6 ions were accumulated) from 350 to 1500 m/z. The top 10 intense ions with charge states ≥ 2 were sequentially isolated to 154 a target value of 1E5 (maximum injection time of 110 ms), fragmented by HCD (NCE 28%) and detected in the Orbitrap at R = 15,000, m/z 200. Data analysis Identification and isotopic quantification of proteins were performed on raw output files from LC-ESI-MS/MS using MaxQuant (Version 1.5.8.3) (Cox et al., 2011) together with its built-in search engine Andromeda. Carbamidomethylation of cysteines was set as a fixed modification, acetylation of protein N-termini, methionine oxidation was included as variable modifications. For quantitative analysis, dimethyl labeled DimethlyLys0, DimethylNter0, DimethlyLys4, DimethylNter4, DimethlyLys8 and DimethylNter8 were used as labels together with the optional iBAQ (Intensity Based Absolute Quantification) calculation. Parent mass tolerance was set to 4.5 ppm (after refinement by MaxQuant) and fragment mass tolerance to 20 ppm. Trypsin was set as the digestion enzyme with up to two missed cleavages allowed. The match between runs feature of MaxQuant was used to transfer peptide identifications from one run to another based on retention time and mass to charge ratio. Both peptide and protein identifications were reported at a false discovery rate (FDR) of 1%. Maxquant was used for quantitation of labelled peptides and the calculation of normalized protein ratio. Phylogenetic analysis Amino acid sequences for DGS1 or NCA2 related proteins were obtained by blast searches using either the yeast NCA2 or Arabidopsis DGS1 against the species listed in Supplemental Dataset 4A (Altschul et al., 1990). Full-length amino acid sequences were aligned using Clustal Omega (Li et al., 2015) and gaps were removed using TrimAI with 155 the gappyout algorithm (Capella-Gutierrez et al., 2009) (Supplemental Dataset 4B). The resulting sequence alignment was used to create a maximum-likelihood phylogeny using the program IQTREE (Nguyen et al., 2015). Branch support values were gained by bootstrapping with 1000 replicates. Co-expression analysis The 200 top co-expressed genes were obtained from the Genevestigator Co-expression tool using the perturbation transcriptomic dataset (10615 samples for Arabidopsis and 1771 samples for S. cerevisiae). All pairwise Pearson’s correlations coefficients were calculated in R and the network was visualized in Cytoscape v3.5.1. Edges were displayed if the Pearson correlation coefficient (PCC) was higher than 0.65 (DGS1 network) or 0.75 (NCA2 network). Accession Numbers DGS1 (At5g12290), MIC60 (At4g39690), TOM40 (At3g20000), Tom20-2 (At1g27390), Tom20-3 (At3g27080), Tom20-4 (At5g40930), RISP (At5g13430) Supplemental Data Sets The additional data sets may be found at: https://doi.org/10.1105/tpc.18.00885. Supplemental Dataset 1. Mass Spectrometry of Supercomplex I+III, complex I, complex V, complex III and the F1 sub-complex of complex V excised from BN-PAGE. Supplemental Dataset 2. Mutant and complementation lines used in the study. 156 Supplemental Dataset 3. Hierarchical clustering of 720 genes differentially expressed (|log2 FC| > 1 and FDR < 0.05) in dgs1-1 and dgs1-1 Comp (L) compared to wild type under drought stress conditions. Supplemental Dataset 4. A) List of organisms and protein accession numbers where a DGS1/NCA2 type ortholog could be detected. B) Aligned amino acid sequences for DGS1 or NCA2 related proteins with gaps removed as outlined in Methods. Supplemental Dataset 5. List of the top co-expressed genes for A) DGS1 and B) NCA2. Supplemental Dataset 6. Primers used in this study. Supplemental Dataset 7. List of antibodies used in this study. Acknowledgment We would like to thank Asha Haslem for technical assistance and the La Trobe University Genomics Platform for access to next generation sequencing equipment. Mass spectrometry was undertaken at the Proteomic Platform, La Trobe University. Electron microscopy was undertaken at the Biosciences Microscopy Unit, School of Biosciences, the University of Melbourne and the RMIT Microscopy & Microanalysis Facility, RMIT University. Confocal microscopy was undertaken at LIMS Bioimaging facility, La Trobe University. We thank Juliette Jouhet for providing the MIC60 antibody. This work was supported by the Australian Research Council Centre of Excellence in Plant Energy Biology (CE140100008) to JW. IDC is supported by the Fonds Wetenschappelijk Onderzoek (Postdoctoral Fellowship no. 12N2415N and Travel Grant for a long stay abroad no. 450215N). JS is supported by a Feodor Lynen Research Fellowship (Alexander von Humboldt Foundation, Germany). The contribution by AL and 157 CB was supported by the Division of Chemical Sciences, Geosciences and Biosciences, Office of Basic Energy Sciences of the United States Department of Energy (DE-FG02- 91ER20021), MSU AgBioResearch, and partially supported by a fellowship from Michigan State University under the Training Program in Plant Biotechnology for Health and Sustainability (T32-GM110523). Author Contribution YW, IDC and JW conceived the project. LL performed the experiments with XM contributing with physiological analyses. OB carried out the RNA-Seq and IDC analyzed the RNA-Seq data and performed the co-expression analyses. AvdM carried out the electron microscopy. JS measured the activities of respiratory chain complexes and quantified the H2O2 content together with YW. AL and CB carried out and interpreted the lipid analysis. CC carried out the phylogenetic analysis. LL drafted the manuscript that was edited by JW, IDC and YW. The final version was produced with contributions from all authors. 158 APPENDICES 159 APPENDIX 5A Main Text Figures 160 Figure 5.1. DGS1 is present in a large multi-subunit protein complex with MIC60, TOM40, TOM20s and RISP. A) Immunodetection of DGS1, MIC60, TOM40, complex III subunit RISP, and complex IV subunit cytochrome oxidase II (COXII) in total mitochondrial proteins separated by BN-PAGE. Coomassie blue staining was performed showing the distribution of supercomplex and complex I to V. MW, molecular weight. B) Mitochondrial proteins from wild-type (Col-0) plants were incubated without or with antibodies raised against DGS1 and MIC60. The wash and protein A-agarose pellet fractions were resolved by SDS-PAGE and immunodetected with antibodies as shown. The interaction between proteins is indicated by asterisks and the corresponding molecular weight for each protein is indicated in kDa. C) Mitochondrial proteins incubated with or without cross-linker were resolved by SDS-PAGE and followed by immunodetection. Red lines indicate proteins that exist in the same complex with DGS1 while blue lines indicate association with another complex. The size of non-crosslinked protein is indicated in each panel. 161 Figure 5.2. A single point mutation in DGS1 alters the multi-subunit complex. A) A schematic gene (left panel) and protein (right panel) model of DGS1. The position of the 162 Figure 5.2 (cont’d) EMS point mutation and T-DNA insertion is indicated. Primers used for screening of homozygous plants are indicated as LP, RP and LB (Supplemental Dataset 6). Two transmembrane domains, the Nuclear Control of ATPase 2 (NCA2) domain and the fragment used for the generation of the DGS1 antibody are indicated in different colors. B) The protein abundance of DGS1, MIC60, TOM20s, TOM40, RISP and COXII was determined by immunoblot analysis of mitochondrial proteins isolated from wild-type (Col-0), dgs1-1 and dgs1-2 mutant lines as well as several lines expressing the dgs1-1 mutant protein. Ten micrograms and twenty micrograms of mitochondrial proteins were loaded to ensure linearity of detection. Mean values are shown for three biological replicates. P-value ≤ 0.05 from a Student’s t-test is indicated with red asterisks. C) Freshly isolated mitochondria (Mit) and outer membrane ruptured mitochondria (Mit*OM) from wild-type (Col-0), dgs1-1 and dgs1-2 mutant lines as well as several lines expressing the dgs1-1 mutant protein were treated with protease K at different concentrations, followed by immunodetection. Proteins displaying altered protease K sensitivity are indicated with red asterisks. D) Crosslinked and non- crosslinked mitochondrial proteins from wild-type (Col-0) plants and dgs1-1 Comp (L) plants were resolved by SDS-PAGE followed by immunodetection. Red asterisks indicate the protein abundance in the crosslinked complex was reduced in the dgs1-1 Comp (L) line, while the blue asterisk indicates the protein abundance was increased. The size of non-crosslinked protein is indicated in each panel. 163 Figure 5.3. The dgs1-1 mutation alters mitochondrial size. A) Protoplasts isolated from the indicated genotypes were transiently transformed with AOX-RFP (mitochondria targeting construct) and KDEL-YFP (ER targeting construct), respectively. Chloroplasts were observed using chlorophyll auto-fluorescence. Scale bars = 10 μm. The rectangle areas were zoomed in to show the altered mitochondrial and chloroplast size; scale bars = 5 μm. The diameter of chloroplasts and mitochondria were averaged from 30 protoplast 164 Figure 5.3 (cont’d) cells and the significant differences were assessed using Students’ t-test with P ≤ 0.05 indicated by asterisks. The white line(s) indicate the diameter measurement used. B) Electron microscopy images of mitochondria cross sections from leaf tissue of the Col-0, dgs1-1 line, dgs1-1 Comp (L) line and dgs1-1 Comp (H1) line. The internal cristae membranes are indicated with white arrows. Scale bars = 0.2 μm. 165 Figure 5.4. The dgs1-1 mutation alters mitochondrial lipid composition. Lipid analysis by Gas-Liquid Chromatography (GLC) of chloroplasts (A) and mitochondria (B) extracted from wild-type (Col-0) and dgs1 mutant lines. MGDG, 166 PC, phosphatidylcholine; sulfoquinovosyldiacylglycerol; Figure 5.4 (cont’d) monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; SQDG, PG, phosphatidylglycerol; PE, phosphatidylethanolamine; CL, cardiolipin. Significant differences in relative composition of each acyl chain and total lipids were determined by Students’ t-test with P-value ≤ 0.05 (n = 3) indicated by asterisks, with error bars = SE. C) The construct expressing full-length DGS1 protein or dgs1-1 mutant protein with C- or N- terminal GFP was transiently transformed into protoplasts isolated from Arabidopsis Col-0 plants. SSU-RFP (targeting to chloroplasts), AOX-RFP (targeting to mitochondria) or KDEL-RFP (targeting to ER) were co-transformed as controls. Scale bars = 10 μm. D) Proteins of chloroplasts, mitochondria and ER isolated from Col-0 and dgs1 mutant plants were separated by SDS-PAGE and followed by immunoblotting using the DGS1 antibody, the TOC86 antibody as a chloroplast marker, the TIM17-2 antibody as a mitochondria marker, and BiP as an ER marker. 167 Figure 5.5. The dgs1-1 mutation affects mitochondrial protein abundance, protein import and alternative respiratory capacity. A) The protein abundance of various 168 Figure 5.5 (cont’d) mitochondrial protein import components and respiratory chain components was determined by immunoblot analysis of mitochondria isolated from Col- 0 and dgs1 mutant plants. 10 µg and 20 µg of mitochondrial proteins were loaded for linearity of detection. Mean values are shown for three biological replicates. P-value ≤ 0.05 from a Student’s t-test is indicated with red asterisks. B) The activity of mitochondrial respiratory chain complexes from wild-type (Col-0), mutant and complementation lines was measured using a Clark-type oxygen electrode and are shown as means ± SE of three biological replicates. Asterisks indicate the significant differences with * P < 0.05; ** P < 0.01 and *** P < 0.001 between oxygen consumption of wild-type mitochondria (Col- 0) and mitochondria from different dgs1 mutants as determined by two-way analysis of variance (ANOVA) with a post-hoc Tukey HSD test. C) In vitro protein import of [35S]-Met- radiolabeled AOX and ANT into mitochondria isolated from Col-0 and dgs1-1 Comp (L) plants. Aliquots were removed at 5, 10 and 15 min and then treated with protease K. The left panel shows a typical image of an in vitro import assay, the precursor and mature (m) forms of the protein are indicated. The right panel shows the rate of import was determined at all time points and normalized to Col-0 at the last time point for each replicate (n ≥ 3). Standard errors for average ratios are indicated on graphs. Asterisks indicate the significant differences compared to Col-0 (P < 0.05, Student’s t-test). 169 Figure 5.6. The dgs1-1 mutation imparts higher drought stress tolerance. A) Wild- type (Col-0) and dgs1 mutant lines were grown under normal conditions for 24 days, then water was withheld for 14 days followed by rewatering for 3 days to recover. Representative plants were imaged at the indicated time points. True leaves 5, 6 and 7 were harvested at the indicated time points and stained for H2O2 (3,3’-diaminobenzidine [DAB]) and O2•− (nitroblue tetrazolium [NBT]). The bar chart shows the survival ratio after 3 days of rewatering. B) The relative water content, maximum quantum yield (Fv/Fm), total chlorophyll content and photosynthesis rate were measured during the drought treatment at the days as indicated. Shown are means ± SE of five biological replicates. Asterisks indicate a significant difference compared to wild-type (Col-0) at each time point (P ≤0.05, Student’s t-test). 170 Figure 5.7. Transcriptome analysis of dgs1 mutants. A) The number of up-regulated genes and down-regulated genes (|log2 (fold change)| >1, FDR<0.05) of each genotype leaves from plants of 24 days old plus 8-day drought treatment, compared to plants grown under normal conditions. B) Heatmap visualization of genes differentially expressed (|log2 (fold change)| >1, FDR<0.05) in dgs1-1 and dgs1-1 Comp (L) compared to wild type during drought stress conditions. Hierarchical clustering of the z-scores of log2- 171 Figure 5.7 (cont’d) transformed transcripts per million values based on Euclidian distance with Genesis 1.6.0 identified three clusters (Blue, Green and Red). C) GO enrichment analysis of genes differentially expressed in dgs1-1 and dgs1-1 Comp (L) during drought stress conditions using ClueGO v2.5.1. The size of the nodes is proportional to the Bonferroni-corrected P value. GO terms were grouped based on their similarity (edge thickness represents the kappa score of similarity between two GO terms) and the most significant term in each group is shown in bold. GO terms were defined as specific for one of the clusters (Blue, Green or Red) if the proportion of associated genes from this cluster is higher than 50%. Common GO terms are indicated in grey. 172 Figure 5.8. Phylogenetic analysis of DGS1- and NCA2-related proteins. A) A maximum-likelihood phylogenetic tree of DGS1- and NCA2-related proteins. Numbers represent ultrafast bootstrap values after 1000 replicates from IQTREE. Some numbers have been manually moved for better visibility. B) DGS1 and NCA2 co-expression 173 Figure 5.8 (cont’d) networks displaying the 200 top co-expressed genes. C) A model of the DGS1 and dgs1-1 mutant protein in plant mitochondria. For the native DGS1 protein, an ability to pull down TOM40, MIC60 and RISP along with cross-linking results suggests a direct interaction. The association with TOM20 is not defined and may be via TOM40. When dgs1-1 mutant protein was expressed, MIC60 was more protected by the inner membrane while RISP was more exposed to the intermembrane space. The abundance of TOM40 and MIC60 was reduced in the MICOS, and the abundance of RISP was increased. The alterations of MICOS due to the presence of the dgs1-1 mutant protein may change the contact between the outer and inner membrane. The asterisk indicates the mutation site of the dgs1-1 protein. RISP, the Rieske FeS protein; MICOS, the mitochondrial contact site and cristae organizing system; Cyt c, cytochrome c; MIC60, 60 kDa subunit of MICOS; TOM40, the translocase of outer mitochondrial membrane 40 kDa subunit; TOM20, the translocase of outer mitochondrial membrane 40 kDa subunit. 174 APPENDIX 5B Supplementary Figures 175 Figure 5.S1. The intramitochondrial localization of TOM20-2 and TOM40 in Col-0 and dgs1 mutant lines. Supports Figure 5.2C. Freshly isolated mitochondria (Mit) and outer membrane ruptured mitochondria (Mit*OM) from wild-type (Col-0) and dgs1 mutant lines were followed by immunoblotting of TOM20-2 and TOM40. treated with Proteinase K of different concentrations, 176 Figure 5.S2. Related to Figure 4A and 4B. Leaf lipid analysis of wild type and mutant lines. A) Total Leaf lipids analyzed by Gas-Liquid Chromatography (GLC) total leaf Lipids extracted from wild-type (Col-0) and dgs1 mutant lines, n = 2 with error bars = SE. MGDG, 177 sulfoquinovosyldiacylglycerol; Figure 5.S2 (cont’d) monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; SQDG, PG, phosphatidylglycerol; PE, phosphatidylethanolamine. B) TLC image of mitochondrial lipids is attached with labels, the line with the highest amount of DGS1 point mutant protein has the oligogalactolipids. PC, phosphatidylcholine; 178 Figure 5.S3. The defective growth phenotype of mutants producing the DGS1-1 protein. Supports Figure 6A. A) Developmental progression of Col-0, dgs1-2, dgs1-1, DGS1 Comp and dgs1-1 complemented lines grown under control condition was recorded. 20 individual plants for each genotype were grown and representative plants and detached rosette leaves are presented. Measurements of Fv/Fm and total 179 Figure 5.S3 (cont’d) chlorophyII content at time point as indicated. Averages were calculated based on three biological replicates for each measurement and significant differences are indicated by asterisks (P≤0.05, Student’s t-test). B) Soil-based growth progression was recorded as described in Boyes et al. (2001): stage 1.04, 1.07, 1.10 and 1.14, maximum radius of 4, 7, 10 and 14 rosette leaves >1 mm; stage 5.10, first flower bud visible; stage 6.10, first flower opens and stage 6.9, flowering complete. Representative growth parameters inflorescence height (mm), number of rosette leaves more than 1 mm and maximum rosette radius (mm) across the growth period were measured. Averages and SE were calculated based on 20 plants with asterisks indicating the significant difference using a Student’s t-test (P ≤0.05). C) Root length analysis of Col- 0, dgs1 mutant lines grown on B5 media plate containing 2% (w/v) sucrose at 9 and 16 days after sowing. 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Cellular metabolism demands the maintenance of membranes not only for the chemical composition of the membrane lipid bilayer, but also for membrane biophysical properties like curvature. The thylakoid membranes in the chloroplast, have a unique galactolipid composition, predominately monogalactosyldiacylglycerol (MGDG) and digalactosyldiacylglycerol (DGDG). The highly curved grana stacks of the thylakoids house the photosynthetic machinery. Proteins embedded in and associated with the membrane often depend on the lipid environment to perform their biochemical roles as well as interact with other proteins and substrates. To meet the cellular needs, there must be tight regulation of lipid synthesis and turnover. Some of this regulation occurs at the transcriptional level, where the transcript levels dictate the amount of protein produced for lipid biosynthesis. Transcription itself can be regulated through the action of transcription factors that respond to stimuli from within the cell as well as outside the cell. Some transcription factors, such as WRINKLED1, have already been identified as regulators of basal lipid metabolism. There are also genes that get transcribed due to stress signals, such as those encoding the PLIP lipases. In addition to transcriptional regulation, it seems that there must be post-translational regulation of the lipid biosynthetic enzymes. Role of Proteolysis in Lipid Homeostasis The characterization of the lipid phenotype exhibited by knock-out mutants of RBL10, a rhomboid-like protease, introduces the potential for regulated intramembrane proteolysis 191 (RIP) as a means to modulate lipid metabolism in Arabidopsis. In mammals, RIP has been shown to be involved in lipid homeostasis through processing of the amyloid precursor protein (APP), which is implicated in Alzheimer’s Disease (AD). Cleavage of APP can be controlled by levels of cholesterol and other lipids in the cell, at the same time, lipid homeostasis also seems to be controlled by the APP cleavage products (Grimm et al., 2012a, Grimm et al., 2012b). Another example of RIP controlled lipid metabolism comes from cholesterol feedback regulation, where the sterol regulatory element binding protein (SREBP) is dually processed by site-1-protease and site-2-protease respectively, freeing SREBP from the membrane of the ER under low cholesterol levels (Brown and Goldstein, 2009). As described in Chapter 3, it seems that the presence of RBL10 enables the flipping of phosphatidic acid (PA) across the inner envelope membrane, thereby allowing the biosynthesis of galactolipids such as MGDG and DGDG from chloroplast- derived precursors. The identity of the protein responsible for the translocation of PA is not yet known and adds another facet to the multistep lipid biosynthetic pathway in plastids. Searching for this protein presents a big challenge as it may or may not be the direct protein substrate for RBL10. Rhomboids have been shown to activate signaling ligands (Lee et al., 2001, Urban et al., 2002), raising the possibility that additional protein participants coordinate the expression, localization, or activation of this PA transporter. One traditional approach to identifying participants is to perform a suppressor screen in the rbl10 mutant background. However, since RBL10 seems to have an activating role in a sense it is unlikely that removing downstream participants will rescue the lipid phenotype. However, in the case that the regulatory mechanism is transcriptional repression of the gene encoding the PA transporter, a suppressor screen could reveal 192 such a participant through a rescued lipid phenotype. Screening for recovered MGDG acyl composition would require the use of gas chromatography (GC) lipid analysis (Wang and Benning, 2011), but since the phenotype of rbl10-1 can be seen in the total acyl composition this would remove the need for the use of thin layer chromatography and enable direct analysis of leaf tissue, expediting screening. Aside from searching for loss- of-function (LOF) mutants, it’s possible to discover a gain-of-function (GOF) mutant as there is an increasing number of examples of GOF in the literature (Li et al., 2019). In the event that LOF or GOF mutants cannot be isolated, analyzing changes in transcript abundance between WT and rbl10 mutant would be an approach which could reveal downstream signaling activated by RBL10 proteolytic activity. RBL10 and its Protein Interactome Chapter 4 describes the discovery of RBL10’s oligomeric state and raises the question of how RBL10 performs its function in the context of a large molecular weight complex. Rhomboids have been implicated in enabling the oligomerization of complexes (Stevenson et al., 2007) and few have been reported to interact with other non-substrate proteins (Jeyaraju et al., 2006, Hwang et al., 2016). However, little is known about rhomboids participating in stable complexes. Identifying the residents of the RBL10 complex would be an important first step in understanding the role of this association. Through co-immunoprecipitation (Co-IP) experiments described in Chapter 4, a list of potential complex denizens has been provided. At first glance, the information currently published about the identified candidates provides little explanation on how they might enable RBL10 to regulate PA trafficking. However, some of the candidates likely are 193 involved in photosynthesis and photosystem repair mechanisms. Another set of proteins is involved in carbohydrate metabolism and ATP synthesis. In addition, some proteins seem to be either involved in protein translation or act as chaperones. Taken together these general functions hint at a potential interconnectivity between them. It has been well established that carbohydrate metabolism and lipid biosynthesis are closely linked and altering the balance of one can influence the other (Zale et al., 2016, Zhai et al., 2017) though the mechanism is not exactly understood. Photosynthetically derived sugars provide the carbon for lipid biosynthesis, and ultimately there must be some feedback regulation mechanism tuning photosynthetic flux. How all these proteins would come to interact with RBL10 or each other is not clear, but it is possible that RBL10 could be mediating the signal from carbohydrate synthesis and altering the balance of lipid biosynthesis through the plastid and ER pathways. The immunoprecipitated proteins could also be substrates for RBL10 cleavage since there are a few with transmembrane domains. It is likely that discovering the direct substrate for an active protease will not be trivial, and a more stringent method, such as biotin proximity labeling BioID (Roux et al., 2013), for detecting protein interactors could be used. Detecting protein substrates that are released from the chloroplast membrane and travel to the nucleus would not be detected with the approach used in Chapter 4, however the BioID approach in whole leaf tissues should capture any proteins that came into close proximity to RBL10. Any protein candidate that is identified by Co-IP or BioID labeling, should be further corroborated through another method such as split-ubiquitin yeast two-hybrid (Obrdlik et al., 2004). 194 Role for Autocatalysis of RBL10 Another discovery described in Chapter 4 is that RBL10 exhibits proteolytic activity towards its own carboxy-terminal domain (CTD). A similar finding has been previously reported for the human Presenilin-associated rhomboid-like (PARL) protein, which cleaves its own amino-terminal domain (NTD) based on phosphorylation status at the NTD (Jeyaraju et al., 2006). This mechanism is a form of regulation of the proteolytic activity of PARL, as the NTD processed form PACT is less active towards its substrate Pink1 (Meissner et al., 2015, Shi and McQuibban, 2017). The self-processing of RBL10 is evidence for its proteolytic activity and also might explain why proteolytic assays have been challenging with purified RBL10 protein. It seems that within the plastid, the intact protein can be detected as well as the cleaved form, however, when RBL10 is heterologously produced in E. coli it immediately undergoes self-processing. The mechanism by which self-processing is triggered or blocked could be due to association with protein interactors in the plastid or post-translational modification. We know that RBL10 is part of a large molecular weight complex; so perhaps the association/dissociation with the complex components coordinates autocatalysis. If the amino acid sequence of RBL10 is analyzed using NetPhos3.1 prediction software (http://www.cbs.dtu.dk/services/NetPhos), 5 phosphorylation sites are predicted at the NTD (serine 49, 81, 88, 114, 119) and 2 closer to the CTD (serine 236, threonine 316) with a confidence above 0.9 (1.0 being the highest likelihood). Though these sites are predicted, it would be instructive to experimentally confirm the phosphorylation status of RBL10 by labeling the protein using 32P-adenosine triphosphate (ATP). Understanding 195 the mechanism which controls the autocatalytic activity of RBL10 would further our understanding of how it can positively affect PA trafficking at the molecular level. In the case of PARL’s self-processing, it was also discovered that the NTD fragment was not degraded after cleavage, but exported from the mitochondria to the nucleus (Sik et al., 2004). In this manner, PARL is able to sense the ATP status of the mitochondrion through phosphorylation and upon reduced ATP levels, send a signal to the nucleus. The discovery that the CTD fragment of RBL10 is present in isolated chloroplasts in addition to the full-length protein begged the question of whether this fragment remains in the chloroplast or is exported outside the plastid. If the fragment remains inside the chloroplast, what is its role? Perhaps, it is simply degraded. It is possible that the autocatalytic cleavage of RBL10 provides a potential mechanism for retrograde signaling from the chloroplast to the nucleus. Identifying the exact cleavage site and amino acid sequence of the CTD fragment would allow a better understanding of its potential role. For example, in the rbl10 mutant, the entire protein is gone and can be complemented by the WT gene. What if only the CTD fragment would be sufficient to complement the mutant lipid phenotype? Conversely, would the protein segment without the cleaved fragment sequence have a different phenotype when used to complement the mutant than just the CTD? These experiments could tease apart the role of each fragment. If the CTD travels to the nucleus, we could in essence capture the signal’s downstream effects, observe any changes in lipid metabolism as well as overall phenotypes. Additionally, understanding the role of proteolysis in RBL10’s molecular mechanism could in part explain why there is embryo lethality when the catalytic residues of RBL10 are mutated to alanine and the rbl10 mutant is complemented with those gene 196 variants. The ability of RBL10 to self-cleave, when it is present in the complex, may be an important step to the function of the chloroplast lipid biosynthetic pathway. On the whole, RBL10 appears to be a multifaceted protein with the potential to relay signals from the chloroplast, affecting changes in lipid metabolism. Understanding the mechanism by which RBL10 acts, may identify a sensing mechanism in the chloroplast, add to our knowledge on retrograde signaling from the plastid, and ultimately further our efforts of understanding lipid regulation in plants. Responses to Stress and Lipid Remodeling Plants utilize phosphate for synthesizing numerous biomolecules and biochemical processes. In most organisms, phospholipids dominate the compositional makeup of membranes. Plants however, due to the abundance of thylakoid membranes in green tissues, reduce their phosphate dependence by using galactolipids for their photosynthetic membranes. As sessile organisms, under low phosphate conditions, plants must re-prioritize the use of phosphate for generating membranes and save them for nucleic acids and other essential macromolecules. Under phosphate starvation, plastid galactolipids are exported to extraplastidic membranes to replace phospholipids (Härtel et al., 2000). Galactolipids come into play again during freezing stress response, when the plastid outer envelope protein SENSITIVE TO FREEZING2 (SFR2) protects the chloroplast from dehydration by transferring additional galactose units onto MGDG, creating oligogalactolipids (Moellering et al., 2010, Moellering and Benning, 2011). In Chapter 5 I discuss how the mitochondrial protein DGS1 influences lipid composition not only in the mitochondria, but also the chloroplast. Initially discovered as a suppressor of 197 the dgd1 mutation (Xu et al., 2008, Moellering and Benning, 2010), DGS1 seems to be a part of the mitochondrial contact site and cristae organizing system (MICOS) complex. While the knock-out mutants of WT DGS1 protein do not seem to change lipid metabolism, the gain-of-function point mutation in dgs1-1 seems to initiate changes in lipid transport between the chloroplast, ER, and mitochondria. Plants expressing the dgs1-1 mutant gene also are more drought tolerant compared to WT plants. It seems that the DGS1 protein could be involved in the response to stress in the mitochondrion, and the MICOS complex is important in proper lipid transport between mitochondria and other organelles. How does a mutant DGS1 protein affect lipid transport? There is the possibility that retrograde signaling from the mitochondrion is the mechanism by which the dgs1-1 overexpressing plants mount a stress response. Additional analysis of the single amino acid change from aspartic acid to asparagine at position 457 near the predicted transmembrane region of DGS1 could give insight on how the change could affect the way DGS1 interacts with components of the MICOS complex. Concluding Remarks Overall, plant cells sense and respond to stress through many mechanisms. Some stresses are detected through organelles like the chloroplast and the mitochondrion. Preliminary evidence for retrograde signals being sent from these compartments, as suggested by findings in the chapters of this dissertation, provides an opportunity to further study the role of the investigated proteins on novel mechanisms for sensing the metabolic status or stress condition of the plant. 198 REFERENCES 199 REFERENCES Brown, M.S. and Goldstein, J.L. (2009) Cholesterol feedback: from Schoenheimer's bottle to Scap's MELADL. J Lipid Res, 50 Suppl, S15-27. 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