Medicine, Faculty of
Medical Genetics, Department of
DSpace
UBCV
Dziurdzik, Samantha Katarzyna
Doctor of Philosophy - PhD
University of British Columbia
Specialized proteins facilitate inter-organelle lipid transfer at membrane contact sites (MCSs), regions where organelles are tethered in close proximity. The VPS13 proteins constitute a unique class of lipid transporters that simultaneously bridge organelles at MCSs and channel phospholipids between them. Of importance, mutations in each of the four human VPS13 (VPS13A-D) genes are associated with distinct neurological disorders, and it is thought that loss of lipid transport at specific MCSs underlies the cause of these diverse diseases. Understanding how VPS13 proteins target membranes and how their MCS recruitment is regulated is therefore crucial for developing a better understanding of the underlying disease pathologies. Here, we uncover a conserved VPS13 membrane targeting mechanism using the Saccharomyces cerevisiae model that has a single Vps13 which localizes to multiple organelles and contact sites. We identify a shared motif in the three known organelle-specific adaptors that recruit Vps13, and establish the necessity of these motifs for interaction. We find all adaptors interact at an uncharacterized six-repeat domain that we named the Vps13 Adaptor Binding (VAB) domain. We demonstrate VAB domain repeats 5–6 are sufficient for adaptor interaction and recruitment supporting a model where adaptors compete for binding to a single pocket. Importantly, modeling a spastic ataxia VPS13D missense mutation in this pocket blocks adaptor interactions, Vps13 recruitment, and further results in a vacuolar protein secretion defect that is not associated with any of the known adaptors. These findings support a conserved adaptor binding role for the VAB domain and suggest the presence of as-yet unidentified adaptors in both yeast and humans. Our studies further identify Fmp27/Hob1 and Ypr117w/Hob2 as putative Vps13 structural homologs localized at endoplasmic reticulum (ER)-plasma membrane (PM) contacts. Recruitment of ER-anchored Fmp27 to PM contacts similarly relies on interactions with a previously uncharacterized protein. Importantly, loss of this PM interactor or of both putative paralogous lipid transporters results in a redistribution of accessible sterols supporting a shared role in lipid homeostasis. Collectively, these findings support the classification of Fmp27 and Hob2 as Vps13-like lipid transporters and suggest adaptor-mediated recruitment is a common membrane targeting strategy for this class of proteins.
MEMBRANE CONTACT SITE TARGETING AND FUNCTIONS OF VPS13 AND RELATED BRIDGING LIPID TRANSPORTERS by Samantha Katarzyna Dziurdzik B.Sc., The University of Waterloo, 2017 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Medical Genetics) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2023 © Samantha Katarzyna Dziurdzik, 2023 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Membrane contact site targeting and functions of Vps13 and related bridging lipid transporters submitted by Samantha Katarzyna Dziurdzik in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Medical Genetics Examining Committee: Elizabeth Conibear, Professor, Medical Genetics, UBC Supervisor Peter Stirling, Associate Professor, Medical Genetics, UBC Supervisory Committee Member Calvin Yip, Associate Professor, Biochemistry and Molecular Biology, UBC University Examiner Robert Molday, Professor, Biochemistry and Molecular Biology, UBC University Examiner Additional Supervisory Committee Members: I. Robert Nabi, Professor, Cellular and Physiological Sciences, UBC Supervisory Committee Member Christopher Loewen, Professor, Cellular and Physiological Sciences, UBC Supervisory Committee Member iii Abstract Specialized proteins facilitate inter-organelle lipid transfer at membrane contact sites (MCSs), regions where organelles are tethered in close proximity. The VPS13 proteins constitute a unique class of lipid transporters that simultaneously bridge organelles at MCSs and channel phospholipids between them. Of importance, mutations in each of the four human VPS13 (VPS13A-D) genes are associated with distinct neurological disorders, and it is thought that loss of lipid transport at specific MCSs underlies the cause of these diverse diseases. Understanding how VPS13 proteins target membranes and how their MCS recruitment is regulated is therefore crucial for developing a better understanding of the underlying disease pathologies. Here, we uncover a conserved VPS13 membrane targeting mechanism using the Saccharomyces cerevisiae model that has a single Vps13 which localizes to multiple organelles and contact sites. We identify a shared motif in the three known organelle-specific adaptors that recruit Vps13, and establish the necessity of these motifs for interaction. We find all adaptors interact at an uncharacterized six-repeat domain that we named the Vps13 Adaptor Binding (VAB) domain. We demonstrate VAB domain repeats 5–6 are sufficient for adaptor interaction and recruitment supporting a model where adaptors compete for binding to a single pocket. Importantly, modeling a spastic ataxia VPS13D missense mutation in this pocket blocks adaptor interactions, Vps13 recruitment, and further results in a vacuolar protein secretion defect that is not associated with any of the known adaptors. These findings support a conserved adaptor binding role for the VAB domain and suggest the presence of as-yet unidentified adaptors in both yeast and humans. Our studies further identify Fmp27/Hob1 and Ypr117w/Hob2 as putative Vps13 structural homologs localized at endoplasmic reticulum (ER)-plasma membrane (PM) contacts. iv Recruitment of ER-anchored Fmp27 to PM contacts similarly relies on interactions with a previously uncharacterized protein. Importantly, loss of this PM interactor or of both putative paralogous lipid transporters results in a redistribution of accessible sterols supporting a shared role in lipid homeostasis. Collectively, these findings support the classification of Fmp27 and Hob2 as Vps13-like lipid transporters and suggest adaptor-mediated recruitment is a common membrane targeting strategy for this class of proteins. v Lay Summary Cells are compartmentalized into organelles that carry out specialized functions necessary for cell survival. Communication between these different compartments occurs at places called “membrane contact sites” where organelles are held closely together. The VPS13 proteins tether organelles together at these sites and channel the fats that make up the compartment walls between them. Mutations in each of the four human VPS13 genes are associated with distinct neurological disorders, and it’s thought that loss of transport between specific subsets of compartments underlies each disease. Using yeast as a model, we identify a shared VPS13 protein region that interacts with compartment-specific adaptors that recruit yeast Vps13 to different cellular sites and further suggest this mechanism is conserved in humans. We additionally identify related Vps13-like proteins with cancer relevance that transport fats between a different set of organelles and find that their recruitment similarly relies on interactions with an adaptor protein. vi Preface A version of Sections 1.5 – 1.7 of Chapter 1 have been published as a review article: Dziurdzik, S.K., and Conibear E. 2021. The Vps13 Family of Lipid Transporters and Its Role at Membrane Contact Sites. International Journal of Molecular Sciences. 2021, Vol. 22, Page 2905. 22:2905. doi:10.3390/IJMS22062905. As first author, I conceived and wrote the manuscript, created all figures and Dr. E. Conibear assisted with revisions. A version of Chapter 2 has been published as a research article: Bean, B.D.M., Dziurdzik, S.K., Kolehmainen, K. L., Fowler, C.M.S, Kwong, W.K., Grad, L.I., Davey, M., Schluter, C., and Conibear, E. 2018. Competitive organelle-specific adaptors recruit Vps13 to membrane contact sites. Journal of Cell Biology. 217:3593–3607. doi:10.1083/jcb.201804111. I was responsible for the co-immunoprecipitation experiments and densitometry analysis presented in this chapter with C. Fowler contributing replicates to Figures 2.1B-C, 2.1D and 2.2A-B. The blots presented for Figures 2.1B and 2.2A are from C. Fowler, while blots for Figure 2.1D are my own. The remaining co-immunoprecipitation experiments in Figures 2.2D-E, 2.3B-C and 2.3D were performed in entirety by me. The original manuscript was conceived and written by Dr. B. Bean and revised by me and Dr. E. Conibear. The domain schematics, sequence alignment and adaptor motifs in Figures 2.1-2.3 were generated by Dr. B. Bean. A modified version of Figure 2.4 was published as: Dziurdzik, S.K., Bean, B.D.M. and Conibear, E. 2018. An interorganellar bidding war: Vps13 localization by competitive organelle-specific adaptors. Contact. 1. doi:10.1177/2515256418814621. As first author, I wrote the associated manuscript and generated these figures with consultation and revisions from Dr. B. Bean and Dr. E. Conibear. vii A version of Chapter 3 has been published as a research article: Dziurdzik, S.K., Bean, B.D.M, Davey, M., and Conibear, E. 2018. 2020. A VPS13D spastic ataxia mutation disrupts the conserved adaptor-binding site in yeast Vps13. Human Molecular Genetics. 29:635–648. doi:10.1093/hmg/ddz318. As first author, I was responsible for writing the manuscript, analyzing the data, generating all figures and performing all experiments with the exception of Figure 3.1E which had replicates performed by Dr. B. Bean, and Figures 3.2C and 3.3E which were performed in entirety by Dr. B. Bean. The analysis of the associated microscopy data and figure generation were performed by me. Dr. E. Conibear and Dr. B. Bean assisted in editing the manuscript. Chapter 4 is original unpublished work performed as part of a collaborative project with a recent graduate from our lab, Dr. V. Sridhar. All experiments performed and figures presented in this chapter were completed by me. The R script used for computing correlation scores presented in Figure 4.11 was generated by O. Riyadh and the original network shown in Figure 4.11D was generated by Dr. E. Conibear. Preliminary observations leading to Figure 4.3A-C and 4.6D were performed by Dr. V. Sridhar. The HHpred analysis in Figure 4.1A-B, and drug spotting in Figure 4.12A were performed concurrently with Dr. V. Sridhar with only my own analysis or replicates presented. I conceived and wrote the associated text in this chapter with revisions provided by Dr. E. Conibear. Chapter 5 is unpublished material that I have written, including a figure I generated, with revisions provided by Dr. E. Conibear. viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables .............................................................................................................................. xiii List of Figures ............................................................................................................................. xiv List of Abbreviations ................................................................................................................. xvi Acknowledgements .................................................................................................................... xxi Dedication ................................................................................................................................. xxiii Chapter 1: Introduction ................................................................................................................1 1.1 Cellular compartmentalization ........................................................................................ 1 1.2 Lipids at organelle membranes ....................................................................................... 1 1.2.1 Glycerophospholipids ............................................................................................. 2 1.2.2 Sphingolipids .......................................................................................................... 4 1.2.3 Sterols ..................................................................................................................... 4 1.2.4 Lipid heterogeneity within organelle bilayers ........................................................ 5 1.3 Intracellular lipid transport ............................................................................................. 7 1.3.1 Vesicular lipid transport and evidence for alternate pathways ............................... 7 1.3.2 Non-vesicular lipid transport at membrane contact sites ........................................ 9 1.4 Lipid transfer protein modules ...................................................................................... 10 1.4.1 Shuttling boxes ..................................................................................................... 11 1.4.2 Sliding tubes ......................................................................................................... 13 ix 1.4.3 Bridging channels ................................................................................................. 14 1.5 VPS13 proteins and disease .......................................................................................... 16 1.6 VPS13 proteins have a variety of localization determinants ........................................ 17 1.6.1 The N-terminus as an ER-targeting determinant .................................................. 19 1.6.2 Organelle-specific adaptors target Vps13 to membranes ..................................... 20 1.6.3 Putative membrane binding APT1 domain ........................................................... 21 1.6.4 Conserved amphipathic helices in ATG2_C domains .......................................... 22 1.6.5 Pleckstrin homology domain ................................................................................ 22 1.7 Function of Vps13 at specific contact sites ................................................................... 23 1.7.1 Vps13 at the NVJ .................................................................................................. 24 1.7.2 Vps13 at mitochondrial contact sites .................................................................... 24 1.7.3 Vps13 and prospore membrane expansion ........................................................... 26 1.7.4 Vps13 at autophagosomal membranes .................................................................. 27 1.7.5 VPS13 proteins at lipid droplets ........................................................................... 29 1.7.6 Vps13 at other contact sites .................................................................................. 30 1.8 Research objectives ....................................................................................................... 30 Chapter 2: Competitive organelle-specific adaptors recruit Vps13 to membrane contact sites ................................................................................................................................................33 2.1 Prelude .......................................................................................................................... 33 2.2 Introduction ................................................................................................................... 34 2.3 Results ........................................................................................................................... 37 2.3.1 The Vps13 DUF1162 domain binds the Ypt35 PxP motif ................................... 37 2.3.2 The DUF1162 domain is part of a 6-repeat region ............................................... 39 x 2.3.3 Spo71 binds the Vps13 DUF1162 domain through a PxP motif .......................... 40 2.3.4 Mcp1 is a mitochondrial PxP motif-containing Vps13 adaptor ............................ 42 2.3.5 Vps13 adaptors compete to recruit Vps13 to different membranes ...................... 45 2.4 Discussion ..................................................................................................................... 45 2.4.1 PxP motif-containing adaptors compete for Vps13 binding ................................. 46 2.4.2 Possible bridging role for Vps13 at contact sites .................................................. 48 2.4.3 The adaptor-binding domain of Vps13 is a conserved 6-repeat region ................ 49 2.5 Materials and methods .................................................................................................. 50 2.5.1 Yeast strains and plasmids .................................................................................... 50 2.5.2 Co-immunoprecipitation ....................................................................................... 52 2.5.3 MEME and FIMO-based motif search ................................................................. 52 Chapter 3: A VPS13D spastic ataxia mutation disrupts the conserved adaptor-binding site in yeast Vps13 ...............................................................................................................................54 3.1 Prelude .......................................................................................................................... 54 3.2 Introduction ................................................................................................................... 55 3.3 Results ........................................................................................................................... 58 3.3.1 Specific VAB domain mutations disrupt binding to the endosomal adaptor ........ 58 3.3.2 Similar VAB domain mutations disrupt the Vps13-Mcp1 interaction ................. 61 3.3.3 All three adaptors have similar requirements for VAB domain binding .............. 62 3.3.4 VAB domain mutations result in CPY secretion .................................................. 63 3.3.5 VAB repeats 5-6 are sufficient for adaptor binding .............................................. 66 3.3.6 Human disease-causing mutations disrupt adaptor binding in yeast Vps13 ......... 68 3.3.7 The VAB domain is comprised of six ß-sandwich folds ...................................... 70 xi 3.4 Discussion ..................................................................................................................... 73 3.4.1 The VAB domain is predicted to form an arc-like structure ................................ 74 3.4.2 New adaptors in yeast and humans ....................................................................... 75 3.4.3 The role of the VAB domain in human VPS13 proteins ...................................... 77 3.5 Materials and methods .................................................................................................. 79 3.5.1 Yeast strains and plasmids .................................................................................... 79 3.5.2 Co-immunoprecipitations ..................................................................................... 81 3.5.3 Fluorescence microscopy and automated quantitation ......................................... 82 3.5.4 CPY secretion ....................................................................................................... 83 3.5.5 Sporulation ............................................................................................................ 83 3.5.6 Statistical analysis ................................................................................................. 84 3.5.7 Protein structure prediction ................................................................................... 84 Chapter 4: A conserved interactor targets Fmp27 to ER-PM contact sites for a shared role in lipid homeostasis ......................................................................................................................85 4.1 Introduction ................................................................................................................... 85 4.2 Results ........................................................................................................................... 88 4.2.1 Fmp27 and Hob2 are RBG proteins with homology to Vps13 ............................. 88 4.2.2 Fmp27 localizes to ER-PM contact sites .............................................................. 91 4.2.3 The uncharacterized protein Ybl086c/Hoi1 recruits Fmp27 to the PM ................ 94 4.2.4 The Hoi1 NT-C2 domain is not sufficient for membrane binding ....................... 97 4.2.5 Recruitment by Hoi1 does not require the Fmp27 C-terminus ........................... 100 4.2.6 ColabFold identifies key interacting residues in Fmp27 .................................... 103 4.2.7 Fmp27 and Hoi1 have a shared role in lipid homeostasis .................................. 108 xii 4.3 Discussion ................................................................................................................... 115 4.3.1 Two conserved Hoi1 helices are required for interaction with Fmp27 .............. 116 4.3.2 Role of Fmp27, Hob2 and Hoi1 in ergosterol homeostasis ................................ 117 4.3.3 Conservation of the lipid homeostasis function of Fmp27 and Hoi1 in higher eukaryotes ........................................................................................................................... 119 4.4 Materials and methods ................................................................................................ 122 4.4.1 Yeast strains and plasmids .................................................................................. 122 4.4.2 Microscopy image acquisition and processing ................................................... 127 4.4.3 Quantitation of microscopic data ........................................................................ 128 4.4.4 Western blot analysis of Fmp27-ENVY and Ybl086c-ENVY ........................... 130 4.4.5 Amphotericin B sensitivity spot assay ................................................................ 131 4.4.6 Protein-protein interaction modeling .................................................................. 131 4.4.7 Chemicogenomic dataset and gene ontology analysis ........................................ 132 Chapter 5: Discussion and Conclusions ...................................................................................133 5.1 Summary of Key Findings .......................................................................................... 133 5.2 Vps13 adaptor proteins bind a single site in the VAB domain ................................... 134 5.3 Additional VAB-interacting adaptor proteins in yeast and humans ........................... 137 5.4 Drivers of lipid transport by bridging channels .......................................................... 141 5.5 The Repeating Beta Groove superfamily .................................................................... 143 5.6 Role of Hob proteins in lipid homeostasis and secretion ............................................ 147 5.7 Limitations and Future Directions .............................................................................. 151 5.8 Concluding Remarks ................................................................................................... 152 Bibliography ...............................................................................................................................154 xiii List of Tables Table 1.1: Localization of human VPS13 proteins ...................................................................... 17 Table 1.2: Localization of yeast Vps13 ....................................................................................... 18 Table 2.1: Yeast strains used in Chapter 2 ................................................................................... 51 Table 2.2: Plasmids used in Chapter 2 ......................................................................................... 51 Table 3.1: Yeast strains used in Chapter 3 ................................................................................... 79 Table 3.2: Plasmids used in Chapter 3 ......................................................................................... 80 Table 4.1: Yeast strains used in Chapter 4 ................................................................................. 123 Table 4.2: Plasmids used in Chapter 4 ....................................................................................... 126 xiv List of Figures Figure 1.1: Membrane lipids and their roles in shaping bilayers. .................................................. 3 Figure 1.2: Modes of intracellular lipid transport .......................................................................... 8 Figure 1.3: Mechanisms of inter-organelle transport by different lipid tranfer modules ............ 12 Figure 1.4: Domain architecture of budding yeast and human VPS13 proteins .......................... 18 Figure 2.1: A central region of Vps13 binds Ypt35 in a PxP motif-dependent manner .............. 38 Figure 2.2: The Vps13 DUF1162 domain recognizes the meiosis-specific adaptor Spo71 through a PxP motif .................................................................................................................................... 41 Figure 2.3: Mcp1 is a mitochondrial PxP motif-containing Vps13 adaptor ................................ 43 Figure 2.4: Model of Vps13 recruitment by adaptor proteins with PxP motifs .......................... 47 Figure 3.1: Mutations in the Vps13 VAB domain disrupt its interaction with the endosomal adaptor Ypt35 ................................................................................................................................ 59 Figure 3.2: VAB domain mutations disrupt Vps13 Mcp1-mediated mitochondrial localization 62 Figure 3.3: VAB domain mutations disrupt Spo71-mediated sporulation and CPY sorting ....... 64 Figure 3.4: VAB domain repeats 5–6 are sufficient for binding all three adaptors ..................... 67 Figure 3.5: Causative Cohen syndrome and spastic ataxia mutations result in perturbed adaptor binding in yeast ............................................................................................................................. 69 Figure 3.6: Causative Cohen syndrome and spastic ataxia mutations result in loss of yeast Vps13 localization and CPY secretion ..................................................................................................... 71 Figure 3.7: AlphaFold modeling of the S. cerevisiae Vps13 VAB domain ................................ 72 Figure 3.8: Model of related membrane recruitment strategies of Vps13 and Atg2 proteins ..... 76 Figure 4.1: HHpred analysis identifies predicted structural homologs of Vps13 ....................... 89 Figure 4.2: Fmp27 resides at ER-PM contact sites ...................................................................... 92 xv Figure 4.3: Uncharacterized protein Hoi1 is the Fmp27 interactor at the PM ............................. 95 Figure 4.4: The PM localization of Fmp27 and Hoi1 is interdependent ..................................... 96 Figure 4.5: The Hoi1 C-terminus is recruited to the ER by Fmp27. ........................................... 98 Figure 4.6: The Hoi1 C-terminus is sufficient for Fmp27 recruitment but the NT-C2 domain alone is not sufficient for membrane targeting ............................................................................. 99 Figure 4.7: Recruitment by Hoi1 does not require the Fmp27 C-terminus ............................... 101 Figure 4.8: ColabFold identifies key interacting residues in Fmp27 ......................................... 104 Figure 4.9: Confidence measures of the Fmp27-Hoi1 ColabFold models ................................ 106 Figure 4.10: The Fmp27 ⍺-helical extension interacts with Hoi1. ........................................... 107 Figure 4.11: Fmp27 and Hoi1 are functionally correlated to ER-PM sterol transporters. ....... 109 Figure 4.12: Fmp27 and Hoi1 have a shared role in lipid homeostasis ..................................... 112 Figure 5.1: RBG lipid transporters in budding yeast ................................................................. 144 xvi List of Abbreviations aECM apical extracellular matrix AH amphipathic helix AmB amphotericin B APT1 aberrant pollen transmission 1 ATG2 autophagy-related 2 ATG2_C autophagy-related protein C-terminal BPI bacterial permeability-inducing protein CETP cholesterol ester transfer protein co-IP co-immunoprecipitation CPY carboxypeptidase Y D4H modified domain 4 D4 of the C. perfringens theta-toxin DAG diacylglycerol DEEPN dynamic enrichment for evaluation of protein networks DHFR dihydrofolate reductase DSF differential scanning fluorimetry DSLS differential static light scattering DUF1162 domain of unknown function 1162 E-SYT extended synaptotagmin EM electron microscopy ER endoplasmic reticulum ERMES endoplasmic reticulum-mitochondria encounter structure xvii FFAT two phenylalanines in an acidic tract FIMO Find Individual Motif Occurrences GalNAc N-acetylgalactosamine GFP green fluorescent protein GO gene ontology GSL glycosphingolipid HA hemagglutinin Hob Hobbit HOI1 hobbit interactor 1 HOP homozygous deletion profiling HOPS homotypic fusion and vacuole protein sorting HRP horse radish peroxidase IMM inner mitochondrial membrane IP-MS immunoprecipitation followed by mass spectrometry IPC inositol phosphorylceramide LAM Lipid transfer proteins Anchored at Membrane contact sites LC/MS/MS liquid chromatography-tandem mass spectrometry LD lipid droplet LPS lipopolysaccharide Ltp lipopolysaccharide transport complex LTPs lipid transfer proteins MCS membrane contact site xviii MEME Multiple Expectation Maximization Algorithms for Motif Elucidation MIPC mannosyl-inositol phosphorylceramide MYLO markerless yeast localization and overexpression NG NeonGreen NT-C2 N-terminal C2 NVJ nucleus-vacuole junction ORD OSBP-related domain ORP Oxysterol-binding Related Protein OSBP OxySterol Binding Protein OSH Oxysterol homology PA phosphatidic acid PAE predicted aligned error pLDDT predicted Local Distance Difference Test PC phosphatidylcholine PCA protein complementation assay PE phosphatidylethanolamine PH Pleckstrin homology PI phosphatidylinositol PI(3,5)P2 phosphatidylinositol 3,5-biphosphate PI(3)P phosphatidylinositol 3-phosphate PI(4,5)P2 phosphatidylinositol 4,5-biphosphate PI(4)P phosphatidylinositol 4-phosphate xix PI(5)P phosphatidylinositol 5-phosphate Pin pharynx ingressed PIP phosphatidylinositol phosphate PM plasma membrane PS phosphatidylserine PTM post-translational modification PX Phox homology PxP proline-X-proline RANK receptor activator of NF-Œ∫B RBG repeating beta groove RFP red fluorescent protein SD synthetic minimal media SHR SHORT-ROOT SHR_BD SHORT-ROOT binding domain SM sphingomyelin SMP synaptotagmin-like mitochondrial-lipid-binding protein StARkin StART-like relatives StART Steroidogenic Acute Regulatory-lipid related Transfer STRIC sterol rich compartment TAG triacylglycerol TAP-MS tandem-affinity purification and mass spectrometry TULIP Tubular lipid binding protein xx UBA ubiquitin-associated VAB Vps13 Adaptor Binding VAMP vesicle-associated membrane protein VancE vacuolar non-NVJ cytoplasmic ER contact site VAPs VAMP-associated proteins vCLAMP vacuole-mitochondria patch VPS13 vacuolar protein sorting 13 WT wild type Y2H yeast 2-hybrid YFP yellow fluorescent protein YPD yeast peptone dextrose media xxi Acknowledgements First and foremost, I would like to express my gratitude to my supervisor, Dr. Elizabeth Conibear, for her guidance and support throughout my studies. I am thankful for all her feedback that has helped shape the direction of my research, and for the opportunities provided that have contributed to my growth as a scientist. I am also sincerely grateful to my committee members Dr. Christopher Loewen, Dr. Peter Stirling and Dr. Robert Nabi for their insightful feedback and guidance during my thesis research. I am fortunate to have had the opportunity to collaborate with several research groups both internationally and within the University of British Columbia. My heartfelt thanks go to Dr. Maya Schuldiner, Dr. Christopher Stefan, Dr. Tim Levine, Dr. Stefan Taubert and Dr. Arash Bashirullah, as well as their talented teams, for the opportunity to collaborate and exchange ideas. These experiences allowed me to explore new areas of research and have been incredibly enriching and inspiring. I would like to express my sincere appreciation to the Canadian Institutes of Health Research Foundation, the BC Children’s Hospital Foundation, and the University of British Columbia for funding my research and enabling me to pursue my passion. My gratitude also goes to the Conibear lab members, past and present, who have made great collaborators and friends. In particular, I would like to thank Mike Davey for his exceptional molecular biology and yeast expertise and for his constant production of yeast strains and plasmids for my research. I am also grateful to Dr. Björn Bean for his mentorship and guidance, to Alexis Shih and Dr. Shawn Whitfield for their training, and to Dr. Vaishnavi Sridhar, Shawn Shortill and Kevin Jeffers for their collaborations. I would also like to thank xxii Sydney Holme whose expertise with mammalian cell culture has been invaluable, and Omer Riyadh and Mia Frier for their thought-provoking discussions and camaraderie. I would also like to thank the students and staff within the department of Medical Genetics and beyond who I have had the pleasure of meeting and working alongside during my degree. Being surrounded by such talented and passionate individuals has been a source of inspiration and motivation throughout my studies. Finally, I would like to extend my deepest appreciation to my family and friends for their patience and encouragement. To my parents and brothers, Robert, Peter and Tomasz, thank you for the love and support. Special thanks go to Guillermo Caballero Silva for his helpful discussions and clever ideas, along with his unwavering support and belief in my abilities. xxiii Dedication Dla moich rodziców i braci 1 Chapter 1: Introduction 1.1 Cellular compartmentalization Eukaryotic cells are compartmentalized into organelles that carry out specialized tasks. Compartmentalization is needed to efficiently coordinate organelle functions by separating incompatible processes and physically grouping related molecules together (Alberts et al., 2002a; Zecchin et al., 2015; Bowsher and Tobin, 2001). Membranes form the physical barriers that enclose organelle compartments, separating their lumen from the cytosol. The main biological component of membranes are lipids, which form fluid bilayers at organelle membranes (van Meer et al., 2008; Alberts et al., 2002a; b). Although compartmentalization supports specialized organelle functions, it necessitates coordinated inter-organelle communication for cells to respond globally to environmental or internal cues. One place that highlights this type of communication is at membrane contact sites (MCSs), regions where organelles are held closely together (Helle et al., 2013; Scorrano et al., 2019; Prinz et al., 2020). MCSs are critical regions of lipid transport, which will be discussed further in this chapter. First, the roles of membrane lipids as both physical barriers and modifiers of membrane properties will be outlined. Further, intracellular lipid transport pathways will be discussed, particularly protein-mediated transport at MCSs with a focus on the membrane-bridging VPS13 lipid transfer proteins. 1.2 Lipids at organelle membranes Membrane lipids have diverse structures and compositions that impart different properties to their associated membranes. Unsurprisingly, different organelles have distinct lipid compositions that provide organelle identity and allow associated proteins to perform their 2 specialized functions (van Meer et al., 2008; van Meer, 2005; Yang et al., 2018). For example, peripheral membrane proteins rely on specific interactions with membrane lipids for appropriate targeting, which may be needed for the further recruitment of additional interactors and regulators (Behnia and Munro, 2005; Dickson and Hille, 2019). Additionally, the conformation of peripheral or transmembrane proteins at organelle membranes can be modified by their association with various lipid species, allowing them to take on functional or inhibitory conformations (Bogdanov et al., 2014; Jelokhani-Niaraki, 2022). The major classes of membrane lipids in eukaryotes which will be discussed in this section are glycerophospholipids, sphingolipids and sterols (Fig 1.1A-D) (van Meer et al., 2008; Alberts et al., 2002b). 1.2.1 Glycerophospholipids Glycerophospholipids are the main structural lipid in eukaryotic membranes (Fig 1.1A,B) (van Meer et al., 2008). Common examples of glycerophospholipids include phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI), and phosphatidic acid (PA). These lipids are comprised of a diacylglycerol (DAG) backbone with two hydrophobic fatty acyl chains and a charged polar headgroup (van Meer et al., 2008; Alberts et al., 2002b). This composition provides glycerophospholipids with an overall cylindrical shape that allows molecules to self-assemble into bilayers where the polar headgroups face the aqueous phase, while the hydrophobic tails are shielded between bilayer leaflets (Fig 1.1E). The packing of glycerophospholipids at membranes is affected by the size and charge of their polar head groups as well as by the length and saturation of their hydrophobic fatty acyl chains (van Meer et al., 2008; Alberts et al., 2002b). Smaller polar headgroups, such as the one in PE, or acyl chain unsaturation which introduce kinks in the hydrophobic tails can confer a 3 Figure 1.1: Membrane lipids and their roles in shaping bilayers. Structures of two common glycerophospholipids: (A) phosphatidylcholine (PC) and (B) phosphatidylethanolamine (PE) with saturated palmitate (C16:0) or cis-unsaturated oleate (C18:1) fatty acyl chains. (C) Structure of sphingomyelin (SM) a common sphingolipid with a characteristic trans-unsaturated sphingosine (C18:0), and an unsaturated (C22:0) fatty acyl chain. (D) Structure of cholesterol, a common sterol. The hydrophobic fatty acyl chains in PC, PE and SM, as well as the bulky hydrophobic region of cholesterol are labelled in orange. (E) Simplified cartoon schematic of a membrane bilayer comprised of conical lipid species such as glycerophospholipids and sphingolipids. Polar headgroups are depicted as blue circles and fatty acyl chains as orange rectangles. (F) Schematic demonstrating how conical lipid species (red or magenta) can introduce curvature into bilayers. (G) Simplified schematic demonstrating how sphingolipids (purple), in association with sterols (yellow), can form liquid ordered domains otherwise known as lipid rafts. more conical lipid structure that can introduce curvature into membranes (Fig 1.1B,F) (Marsh, 2007; van Meer et al., 2008). The introduction of membrane curvature is particularly important 4 at sites of vesicle budding and fusion (Day and Stachowiak, 2020; Mironov et al., 2020). Additionally, fatty acyl chain unsaturation can disrupt tight lipid packing and improve membrane fluidity (Ballweg and Ernst, 2017; Alberts et al., 2002b). Membrane fluidity is required for free lateral movement of lipids and transmembrane proteins to occur within a bilayer. 1.2.2 Sphingolipids Sphingolipids comprise a class of cylindrical membrane lipids that localize almost exclusively to the external leaflet of the plasma membrane (PM) at the cell exterior (Fig 1.1C) (van Meer et al., 2008; Gault et al., 2010). The most abundant sphingolipids differ across eukaryotes with sphingomyelin (SM) and glycosphingolipids (GSLs) predominating in humans (van Meer et al., 2008), while inositol phosphorylceramide (IPC) and mannosyl-inositol phosphorylceramide (MIPC) are more abundant in unicellular yeast (Cowart and Obeid, 2007). Members of this lipid class are comprised of a ceramide backbone and have charged head groups and hydrophobic fatty acyl tails (Gault et al., 2010; Alberts et al., 2002b). Unlike glycerophospholipids, the fatty acyl chains of sphingolipids tend to be longer and saturated. The narrower cylindrical structure and long hydrophobic tails allow sphingolipids to tightly pack at membrane bilayers and, if not mixed with other lipid species, form a rigid gel phase (van Meer, 2005; Alberts et al., 2002b). 1.2.3 Sterols Sterols are bulky non-polar lipids that are essential for membrane fluidity and permeability (Fig 1.1D) (Mesmin and Maxfield, 2009; Liscum and Munn, 1999). Cholesterol is the primary sterol in humans, while ergosterol is the most abundant in yeast (van Meer et al., 5 2008). Sterols co-distribute with lipids within bilayers through weak interactions between the polar hydroxyl head group of sterol and the charged headgroups of sphingolipids or glycerophospholipid species (Maekawa and Fairn, 2015; Guan et al., 2009). Here, the association of the bulky hydrophobic region of a sterol molecule with the fatty acyl chains of nearby lipids result in steric hinderance that can modulate membrane fluidity. This steric hinderance makes sterol-rich membranes less deformable, and thus less permeable (Alberts et al., 2002b). The PM, an organelle that must regulate entry and exit from the cell, has the highest proportion of cellular sterols (~60% total sterol) making for a non-permeable barrier (Mesmin and Maxfield, 2009; Solanko et al., 2018). Sterols at the external leaflet of the PM associate with sphingolipids and introduce fluidity into their rigid bilayers (van Meer et al., 2008; Alberts et al., 2002b). Complexes between sterols and sphingolipids are also important for the formation of liquid ordered domains within membranes, also known as lipid rafts (Fig 1.1G) (Bieberich, 2018; Ripa et al., 2021; Lingwood and Simons, 2010). The tight association of the longer sphingolipid tails result in membrane thickening that can exclude certain transmembrane proteins and induce the recruitment of other peripheral membrane proteins (Lingwood and Simons, 2010; Alberts et al., 2002b). Liquid ordered domains can therefore laterally separate and concentrate specific proteins and effectors together for shared functions, such as in receptor activation and signal transduction (Varshney et al., 2016; Mollinedo and Gajate, 2020). 1.2.4 Lipid heterogeneity within organelle bilayers In addition to variations in lipid composition between organelles, the composition between membrane bilayers is also asymmetric and heterogenous (van Meer et al., 2008; van 6 Meer, 2011). Certain lipid species localize predominantly to luminal/extracellular leaflets such as SM, while others are largely restricted to cytosolic leaflets such as PS (Alberts et al., 2002b; Lorent et al., 2020). This discrete localization is functionally necessary, as is well exemplified by PS whose presence in cytosolic PM leaflets is required for a number of signaling pathways. Conversely in apoptotic cells, PS is exposed to the extracellular PM leaflet where it acts as a potent “eat me” recognition signal that directs immune cells to induce phagocytosis (Kay and Grinstein, 2013; Sebastian et al., 2012; Leventis and Grinstein, 2010; Calianese and Birge, 2020; Kay and Fairn, 2019). Transbilayer phospholipid transport is performed by three types of translocator: flippases, floppases and scamblases (van Meer, 2011; Montigny et al., 2016). Flippases belong to the P4-ATPase family that “flip” specific phospholipid species from luminal/extracellular to cytosolic leaflets in an ATP-dependent manner, while similarly ATP-dependent floppases, such as ABC transporters, conversely “flop” specific lipids from cytosolic to luminal/extracellular leaflets (Pomorski and Menon, 2006; Muthusamy et al., 2009; Montigny et al., 2016). This regulated transport can result in an asymmetric number of lipids in one bilayer versus the other which has been shown to induce membrane curvature and is required for vesicle budding events (Sebastian et al., 2012; Graham, 2004; Xu et al., 2013; Sheetz and Singer, 1974). On the other hand, scramblases equilibrate membranes bilayers by performing non-specific, ATP-independent bidirectional lipid transport (Montigny et al., 2016; Bevers and Williamson, 2010; Williamson, 2016). 7 1.3 Intracellular lipid transport With the variation in organelle membrane lipid compositions, it is striking that the majority of lipid precursors are synthesized at a single organelle – the endoplasmic reticulum (ER) (van Meer et al., 2008). Here, sterols and the precursors for glycerophospholipids (PA) and sphingolipids (ceramide) are synthesized and must be redistributed to their target membranes (Blom et al., 2011; Yang et al., 2018; Balla et al., 2020). These precursors, as well as exogenously supplied lipids, can be further modified by lipid biosynthetic enzymes that are spatially segregated to particular organelle membranes (Balla et al., 2020; Blom et al., 2011). For example, in humans and yeast the decarboxylase that converts PS to PE is found at the double bilayer-containing mitochondria, where it is restricted to the innermost bilayer known as the inner mitochondrial membrane (IMM) (Balla et al., 2020; Tamura et al., 2012; Calzada et al., 2019). Therefore, intracellular lipid transport is critical to ensure the appropriate distribution of lipid species within membranes. 1.3.1 Vesicular lipid transport and evidence for alternate pathways Vesicular trafficking is one method by which intracellular lipid transport can occur. In this process, transport vesicles bud from one organelle membrane and fuse with another (Fig 1.2A) (Bonifacino and Glick, 2004; Alberts et al., 2002c). This is crucial for regulated transport of transmembrane and luminal proteins, which are selectively sorted into vesicles and transported along the endocytic and exocytic pathways (Cui et al., 2022; Alberts et al., 2002c). However, vesicular transport of lipids is much less specific as bulk membranes are transported between compartments. Thus, vesicular transport cannot be sufficient to modify membrane lipid 8 Figure 1.2: Modes of intracellular lipid transport. (A) Cartoon depiction of vesicular trafficking between the ER and Golgi. In this process transport vesicles bud from one organelle, are trafficked through the cytosol and then fuse at another organelle. Membrane lipids undergo non-specific bulk transport while cargo proteins are selectively sorted for transport. (B) Cartoon depiction of non-vesicular lipid transport mediated by lipid transfer proteins at membrane contact sites. Sterol transport at ER-Golgi contact sites mediated by the OSBP-related domain (ORD) of the OSBP lipid transporter is depicted. OSBP interacts with Golgi membranes through its phospholipid-binding Pleckstrin homology (PH) domain and is tethered to the ER by interactions with VAP-A or VAP-B mediated by a FFAT (two phenylalanines in an acidic track) motif. Lipid transport is rapid and specific with sterols being transported to the Golgi, and PI4P transported to the ER (not depicted) (Nakatsu and Kawasaki, 2021). compositions specifically or rapidly in response to cellular cues, nor can it maintain the diverse lipid compositions at different organelles. Other mechanisms were long known to contribute to lipid transport as both chemical and genetic disruptions of vesicular trafficking do not inhibit the transport of various lipids from the ER to the PM (Kaplan and Simoni, 1985; Vance et al., 1991; Baumann et al., 2005; Heino et al., 2000). This is further supported by the more rapid rate of lipid transport between the ER and PM (t1/2 = ~15min), compared to that of vesicle-delivered proteins in related cells (t1/2 = ~60min) 9 (Heino et al., 2000; Baumann et al., 2005). Finally, the presence of organelles that rely on lipid delivery for function but that are not directly connected to vesicular trafficking pathways, such as mitochondria and peroxisomes, provide additional support for the presence of non-vesicular lipid transport pathways (Jain and Holthuis, 2017; Lev, 2010). 1.3.2 Non-vesicular lipid transport at membrane contact sites As previously stated, MCSs are critical regions for intracellular lipid transport. At these sites, organelle membranes are closely tethered together by proteins at an average distance of 15-30 nm (Egea, 2021; Scorrano et al., 2019; Phillips and Voeltz, 2016). The proximity of membranes at these sites enhances the rate and efficiency of lipid transport by allowing specialized lipid transfer proteins (LTPs) to directly move lipid species between membranes (Fig 1.2B) (Jain and Holthuis, 2017; Wong et al., 2019; Levine, 2004). MCSs have been reported for a plethora of organelle pairs, with the lipid-synthesizing ER forming contacts with virtually all organelles (Phillips and Voeltz, 2016; Wu et al., 2018; Prinz et al., 2020; Levine, 2004). The resident proteins at MCSs can function as tethers, regulators or LTPs (see Section 1.4). Protein tethers, which contain transmembrane or membrane-targeting domains, physically bridge organelle membranes at contacts and are necessary to maintain these sites (Eisenberg-Bord et al., 2016; Zaman et al., 2020). Tethers can be comprised of multiple interacting proteins which localize to different organelles, or can be single proteins with multiple organelle-targeting domains. Examples of these include the tether formed at nuclear ER-vacuole contacts through interactions between nuclear ER resident Nvj1 and vacuolar Vac8 in yeast (Jeong et al., 2017; Pan et al., 2000), or tethers formed by the individual extended synaptotagmin (E-SYT) proteins at ER-PM contact sites in humans (Pérez-Lara and Jahn, 2015; Herdman and Moss, 2016; Min et 10 al., 2007). Regulators that affect interactions between tether pairs or that affect the targeting of the membrane-binding domains of tethers can therefore modulate the dynamics of MCSs. Additionally, other regulatory proteins can modify the lipid transfer activity at MCSs by affecting the recruitment or function of their associated LTPs. For example, the vesicle-associated membrane protein (VAMP)-associated proteins (VAPs) are MCS residents found at a number of ER contact sites where they physically interact with a wide variety of LTPs (Murphy and Levine, 2016; Kors et al., 2022; Loewen and Levine, 2005). The interactions of VAP proteins can therefore regulate lipid transfer between the ER and various organelles by enabling transport at specific sites. Other regulators may instead be dispensable for LTP recruitment to contact sites, but are critical for their function. This type of regulation has been observed with the Lipid transfer proteins Anchored at Membrane contact sites (Lam) family in yeast whose sterol transport activity, but not their ER-PM targeting, is affected by interactions with paralogous Laf1/Dgr2 regulators (Topolska et al., 2020). This high level of regulation at MCSs highlights an important role for lipid transport in cell viability. Indeed, disruptions of lipid transport at MCSs are associated with a number of diseases, including neurological disorders and cancer (Nishimura et al., 2004; Datta et al., 2019; Li et al., 2016; Wang et al., 2010; Kim et al., 2022; Wong et al., 2019; Peretti et al., 2020). 1.4 Lipid transfer protein modules LTPs are a large, structurally diverse functional class of proteins (Egea, 2021; Reinisch and Prinz, 2021; Wong et al., 2019). At MCSs, LTPs perform the energetically unfavorable task of transferring hydrophobic lipid species across the aqueous inter-organellar space. The common steps involved in transfer include extraction of lipid molecules from their donor membrane, lipid 11 shielding during transfer, and insertion of lipids into their acceptor membrane (Egea, 2021; Wong et al., 2019; Reinisch and Prinz, 2021). Although lipid transfer domains are structurally variable, many are comprised of highly twisted ß-sheets that form hydrophobic pockets that can accommodate lipid molecules (Wong and Levine, 2017; Reinisch and Prinz, 2021; Drin et al., 2016; Mesmin et al., 2013). Additionally, these modules tend to have membrane bending properties that help with desorption of lipids from donor membranes (Egea, 2021; Wong et al., 2019). The major types of lipid transfer domains and their general mechanisms of transfer will be outlined in this section. 1.4.1 Shuttling boxes One common type of LTP module at MCSs are box-like lipid shuttles that move back and forth between organelles. These modules form box-like hydrophobic lipid binding pockets with flexible lids that can reversibly trap individual lipid moieties and associate with their polar head groups during transfer (Fig 1.3A) (Reinisch and Prinz, 2021; Egea, 2021; Wong et al., 2019). These domains can either freely diffuse between membranes or can be tethered to organelles at contact sites (Egea, 2021; Wong et al., 2019). Several protein families rely on such modules for lipid transport including the Steroidogenic Acute Regulatory-lipid related Transfer (StART) proteins and their StART-like relatives (StARkin), which include the Lam family of yeast sterol transporters (Alpy and Tomasetto, 2014; Horenkamp et al., 2018; Miliara et al., 2019). Other members include the mammalian OxySterol Binding Proteins (OSBPs), Oxysterol-binding Related Proteins (ORPs) and the yeast Oxysterol homology (Osh) proteins which transfer lipids via their OSBP-related domains (ORDs) (Arora et al., 2022; Im et al., 2005; Olkkonen and Li, 2013). 12 Figure 1.3: Mechanisms of inter-organelle transport by different lipid tranfer modules. (A) Lipid transport by shuttling boxes with yeast Osh4 shown as an example. The hydrophobic interior of the Osh4 ORD domain associates with ergosterol or PI(4)P, and functions in counter-current transport of these two lipids at ER-Golgi contact sites. A flexible lid shields bound lipids during transfer across the aqueous cytosol. (B) Crystal structure of the Osh4 ORD domain bound to PI(4)P (PDB 3SPW) or ergosterol (PDB 1ZHZ) shown in yellow. The flexible loop is shown in cyan to highlight the different conformations when bound to the different lipid moieties. (C) Lipid tranport by sliding tubes with human E-SYT2 shown as an example. The E-SYT2 SMP domain homo-dimerizes to form a continuous hydrophobic lipid channel. Lipids are thought to slide into the channels where they can further move between monomers. Lipid transport between the ER and PM is proposed to occur by a shuttling or “pass off” mechanism between different SMP modules. E-SYT2 has a N-terminal hydrophobic stretch (HS) that inserts in the ER. N-terminal C2 domains associate with phospholipids at the ER (C2A and C2B), while the C-terminal C2 domain binds phospholipids at the PM in response to Ca2+ sensing. (D) Crystal structure of dimerized E-SYT2 SMP domains (PDB 4P42) each bound to a PE lipid and Triton X-100 molecule (detergent) labelled in yellow. The different SMP monomers are shown in cyan and purple. (E) Lipid transport by bridging channels with yeast Atg2 shown as an example. The elongated Atg2 protein tethers membanes at either end of the ER-isolation membrane (IM)/autophagosome contact site and directly transports lipids between them through a hydrophobic channel. The segment of the Atg2 channel resolved in the crystal structure in F is indicated by a dashed box. (F) Crystal structure of the N-terminus of the thermophilic yeast, C. thermophilus, Atg2 bound to PE (PDB 6A9J) with the lipid shown in yellow. 13 The ORD domains have been extensively structurally characterized and provide a key example for lipid transfer by box-like shuttles (Tong et al., 2013, 2021; Manik et al., 2017; Im et al., 2005; Moser von Filseck et al., 2015). These proteins are bispecific and transfer different lipid species in opposing directions at contact sites (Arora et al., 2022; Egea, 2021; Wong et al., 2019). Yeast Osh4, which localizes to ER-Golgi contacts, is one example of such a counter-current transporter which traffics phosphatidylinositol 4-phosphate (PI(4)P) to the ER and then efficiently traffic sterols back to the Golgi against their concentration gradient (de Saint-Jean et al., 2011; Koag et al., 2013; Singh et al., 2009). The lid of the Osh4 ORD domain is necessary to accommodate both charged PI(4)P lipids as well as non-polar sterols, and provides a charged region that can flexibly associate with polar lipid head groups (Fig 1.3B). In addition to shielding lipids during transport, the lid has also been found to be necessary for lipid extraction suggesting it may deform donor membranes to aid with lipid desorption (Moser von Filseck et al., 2015). 1.4.2 Sliding tubes Tubular lipid binding proteins (TULIPs) form a superfamily of LTPs with structurally-related lipid transfer modules (Wong and Levine, 2017; Egea, 2021; Wong et al., 2019). These modules from elongated tubular structures with large hydrophobic tunnels that can accommodate multiple lipids, and a lateral seam that associates with polar lipid head groups. These modules have been found to form homo- or hetero-dimers that create a continuous hydrophobic channel and are thought to function by sliding lipids between modules and shuttling between membranes (Fig 1.3C) (Wong and Levine, 2017; Saheki and De Camilli, 2017). TULIP domains are found in the bacterial permeability-inducing protein (BPI), cholesterol ester transfer protein (CETP) and in proteins with synaptotagmin-like mitochondrial-lipid-binding protein (SMP) domains, which 14 include mammalian E-SYTs (tricalbins in yeast) (Wong and Levine, 2017; Wong et al., 2019; Jeyasimman and Saheki, 2020). The first TULIP member to have its LTP domain structurally resolved was the ER-PM glycerophospholipid transporter E-SYT2 (Schauder et al., 2014). The SMP domain of this protein was found to form head-to-head homodimers that create a continuous hydrophobic tunnel that associates with lipid acyl chains, where each monomer binds two lipid molecules (Fig 1.3D). Mass spectrometry analysis of purified lipid-bound E-SYT2 further found that it binds glycerophospholipids with no specificity for lipid head groups due to the open solvent-exposed seam and therefore has less lipid specificity than box-like shuttle LTP domains (Schauder et al., 2014). In support of a lipid sliding and pass-off model for lipid transport, the E-SYT2 SMP dimer is only ~9 nm in length and thus is not a sufficient length to bridge ER-PM contact sites and form a continuous channel (Schauder et al., 2014; Saheki and De Camilli, 2017; Egea, 2021). 1.4.3 Bridging channels The newest identified family of MCS-associated LTPs form elongated lipid transfer channels that physically bridge opposing organelle membranes (Fig 1.3E) (Reinisch and Prinz, 2021; Egea, 2021). These modules are comprised of repeating curved ß-sheets that form an elongated groove with a hydrophobic interior that can accommodate numerous lipid acyl chains, as well as overarching ⍺-helices that can interact with polar lipid head groups (Kumar et al., 2018; Osawa et al., 2019; Reinisch and Prinz, 2021). Unlike the previously described lipid transfer modules, bridging LTPs do not need to diffuse across contact sites and therefore could function in bulk lipid transfer. 15 The large vacuolar protein sorting 13 (VPS13) and autophagy-related 2 (ATG2) proteins with respective amino acid sequences greater than 3,000 and 1,500 residues were the first members of this family to be identified, although others have now been described (John Peter et al., 2022; Toulmay et al., 2022; Neuman et al., 2022b; Hanna et al., 2022). Crystal structures of the N-termini (>300 residues) of C. thermophilum Vps13 (Kumar et al., 2018) and S. pombe Atg2 (Osawa et al., 2019), which encompass their conserved chorein domains, find that they form related structures with an interior hydrophobic groove. The Atg2 N-terminus was co-crystallized with PE confirming that lipid acyl chains contact the hydrophobic groove, however the presence of additional gaps suggests diverse lipid moieties can bind (Fig 1.3F) (Osawa et al., 2019). Indeed, in vitro studies with the equivalent fragment from S. cerevisiae Vps13 determined that this domain can bind multiple glycerophospholipids (Kumar et al., 2018). Further, human ATG2A and N-terminal segments of yeast Vps13 and Atg2 have been shown to be sufficient for glycerophospholipid transport when tethered between liposomes (Kumar et al., 2018; Maeda et al., 2019; Osawa et al., 2019). Several low-resolution negative-stain electron microscopy (EM) or cryo-EM reconstructions of Vps13 and Atg2 proteins have revealed that the full-length proteins form extended structures that are sufficiently long to bridge membranes at contact sites (De et al., 2017; Chowdhury et al., 2018; Valverde et al., 2019). An additional C. thermophilum Vps13 cryo-EM reconstruction confirmed that the N-terminal hydrophobic groove extends far into the protein (Li et al., 2020). In support of a bridging channel model, Vps13 lipid transfer is blocked in a mutant where a restricted ring of hydrophobic residues positioned within the groove was substituted with charged residues. This mutant was found to bind glycerophospholipids but was 16 unable to function in sporulation, a process in which Vps13 is required for prospore membrane expansion and closure (Li et al., 2020). 1.5 VPS13 proteins and disease Prior to uncovering its lipid transfer function, Vps13 was first identified in a screen for yeast mutants defective in sorting carboxypeptidase Y (CPY) to vacuoles (Bankaitis et al., 1986). Yeast Vps13 is now known to be required for a wide range of processes that include transport between Golgi and endosomes, trans-Golgi network homotypic fusion, sporulation, ER macroautophagy and the rescue of mitochondrial defects in mutants lacking the ER-mitochondria tether known as the ER-mitochondria encounter structure (ERMES) (De et al., 2017; Park and Neiman, 2012; Park et al., 2016; Lang et al., 2015; Chen et al., 2020). Humans have 4 homologous VPS13 genes and strikingly, mutations in each are associated with distinct neurological disorders: chorea-acanthocytosis (VPS13A) (Rampoldi et al., 2001), Cohen syndrome (VPS13B) (Kolehmainen et al., 2003), early-onset Parkinson's disease (VPS13C) (Lesage et al., 2016) and ataxia (VPS13D) (Seong et al., 2018; Gauthier et al., 2018). The phenotypic differences associated with mutations in each human VPS13 protein are not explained by differential expression as the homologs are present in all tissues. Instead, the different human VPS13 proteins localize to different sets of MCSs (Table 1.1). The loss of lipid transport at a specific MCS occupied by one VPS13 homolog has been advanced as an explanation for the diverse neurological disorders resulting from mutation of individual VPS13 proteins (Bean et al., 2018; Ugur et al., 2020) 17 Table 1.1: Localization of human VPS13 proteins Homolog Localization Reference VPS13A Endoplasmic reticulum-mitochondria (Kumar et al., 2018; Yeshaw et al., 2019; Muñoz-Braceras et al., 2019) Endoplasmic reticulum-lipid droplet (Kumar et al., 2018; Yeshaw et al., 2019) Mitochondria-endosome (Muñoz-Braceras et al., 2019) VPS13B Golgi apparatus (Seifert et al., 2006) Endosome (Koike and Jahn, 2019) Acrosome (Da Costa et al., 2020) VPS13C Endoplasmic reticulum-endosome/lysosome (Kumar et al., 2018) Endoplasmic reticulum-lipid droplet (Kumar et al., 2018) Phagophore (Rogers and Foster, 2007; Shui et al., 2008; Trost et al., 2009) VPS13D Endoplasmic reticulum-mitochondria (Guillén-Samander et al., 2021) Endoplasmic reticulum-peroxisome (Guillén-Samander et al., 2021) Lipid droplet-mitochondria (Wang et al., 2021) Golgi apparatus (Guillén-Samander et al., 2021) 1.6 VPS13 proteins have a variety of localization determinants Understanding how VPS13 proteins are localized to contact sites where they are thought to function in lipid transport has been a focus of recent research. However, human VPS13 proteins have been challenging to study due to their large size and low expression levels. Yeast has thus proven a useful model organism for identifying the localization determinants and functions of VPS13 proteins due to its single, conserved Vps13 that localizes to multiple organelles and MCSs (Table 1.2). VPS13 homologs from yeast to humans share a number of conserved domains that can contribute to organelle membrane-targeting on either end of the lipid transfer channel (Fig 1.4). In this section, the current understanding of how each conserved domain contributes to membrane targeting will be discussed. However, the role of the Vps13 18 Figure 1.4: Domain architecture of budding yeast and human VPS13 proteins. The shared, conserved domains of VPS13 proteins are labelled. Domains occurring in individual proteins, such as the ubiquitin-associated (UBA) domain of VPS13D, are not shown. Domain abbreviations and their associated Pfam accession IDs are as follows: Chorein (PF12624); VPS13 Adaptor Binding (VAB) (self-defined, see Chapters 2-3); Golgi-body localization domain of maize aberrant pollen transmission 1 (APT1) protein (PF10351); autophagy-related protein C-terminal (ATG2_C) (PF09333); and Pleckstrin homology (PH) (PF00169). ER tethering in VPS13A, VPS13C and VPS13D is achieved through interactions with VAP-A and VAP-B through two phenylalanines (FF) in an acidic tract (FFAT) motifs (Kumar et al., 2018; Guillén-Samander et al., 2021), while the N-terminus of the yeast chorein domain is sufficient for ER targeting (Nakamura et al., 2021). C-terminal VAB, ATG2_C and PH domains are thought to contribute to membrane targeting at the opposing organelle at membrane contact sites. Adaptor Binding (VAB) domain which was identified and characterized as part of the studies in this thesis will only be discussed in Chapters 2 and 3. Table 1.2: Localization of yeast Vps13 Membrane Contact Site Reference Nucleus-vacuole junctions (NVJs) (Park et al., 2016; Lang et al., 2015; Bean et al., 2018) Mitochondria-endosome (Park et al., 2016) Vacuole-mitochondria patch (vCLAMP) (Park et al., 2016; Lang et al., 2015) 19 Single Organelle Reference Endosome (Park and Neiman, 2012; Bean et al., 2018; Rzepnikowska et al., 2017) Mitochondria (Lang et al., 2015; Park et al., 2016; John Peter et al., 2017) Vacuole (Bean et al., 2018; Lang et al., 2015; Park et al., 2016; Chen et al., 2020) Prospore membrane (Park and Neiman, 2012) Golgi apparatus (Kolakowski et al., 2021) Peroxisome (John Peter et al., 2017) 1.6.1 The N-terminus as an ER-targeting determinant A conserved N-terminal helical region within the chorein domain of both Vps13 and Atg2 binds directly to the ER (Kotani et al., 2018a; Nakamura et al., 2021), the major site of lipid biosynthesis (van Meer et al., 2008). This region is highly conserved between both proteins and in fact autophagy defects resulting from loss of ATG2 are rescued by Atg2 chimeras that substitute the N-terminus with that of Vps13 (Osawa et al., 2019), or to a lesser extent the ER-localization signal of Sec71 (Kotani et al., 2018a). Human VPS13A and VPS13C proteins also contain an N-terminal chorein domain, but have a FFAT (two phenylalanines in an acidic tract) motif in a loop adjacent to this region that is required for ER recruitment through interactions with the ER MCS proteins, VAP-A and VAP-B (Kumar et al., 2018; Yeshaw et al., 2019). A non-canonical phospho-FFAT motif, which interacts with VAP proteins only when phosphorylated, was additionally found to target VPS13D to the ER (Guillén-Samander et al., 2021). VPS13B also has a predicted phospho-FFAT but its ER localization has not been studied (Di Mattia et al., 2020). It is unclear if the N-terminus of Vps13 binds the yeast VAP homologs, but so far no predicted FFAT motifs have 20 been found in this region. Therefore, the role of the conserved human chorein domains in membrane targeting, remains an open question. 1.6.2 Organelle-specific adaptors target Vps13 to membranes Yeast Vps13 localizes to multiple organelles and MCSs (see Table 1.2) and binds different membranes under specific nutrient conditions and developmental stages. In vegetative yeast, Vps13 is found primarily at endosomes with a smaller pool at mitochondria (Lang et al., 2015; Park et al., 2016). Under glucose-limiting conditions, Vps13 relocalizes to the ER-vacuole MCS known as the nucleus-vacuole junction (NVJ) (Park et al., 2016; Lang et al., 2015; Bean et al., 2018). In contrast, during sporulation Vps13 is recruited to prospore membranes which form around daughter haploid nuclei (Park and Neiman, 2012). If Vps13 resides at MCSs at these organelles, the bridging model predicts that one end of the lipid transport channel must be anchored to a variety of organelle membranes. Recent work has identified Vps13 adaptor proteins specific to each of the previously described sites that dictate these different localizations. Vps13 is recruited to the prospore membrane by the adaptor Spo71 (Park and Neiman, 2012), to mitochondria by Mcp1 (John Peter et al., 2017) and to endosomes and vacuoles by Ypt35 (Bean et al., 2018). Each adaptor associates directly with an organelle membrane, either through lipid binding PH or Phox homology (PX) domains in the case of Spo71 and Ypt35, or membrane-spanning domains in the case of Mcp1. The mechanism by which these adaptor proteins recruit Vps13 to membranes will be discussed further in Chapters 2 and 3. 21 1.6.3 Putative membrane binding APT1 domain The aberrant pollen transmission 1 (APT1) domain is conserved in both VPS13 and ATG2 proteins and has been shown to interact with specific phospholipids in vitro through phosphatidylinositol phosphate (PIP) strips and liposome binding assays. The APT1 domain in yeast Vps13 binds to phosphatidylinositol 3-phosphate (PI(3)P), while in human VPS13A this domain binds both PI(3)P and phosphatidylinositol 5-phosphate (PI(5)P) (Rzepnikowska et al., 2017; Kolakowski et al., 2020). The APT1 domain of yeast Atg2 has a similar phospholipid preference, binding to PI(3)P, and to a lesser extent phosphatidylinositol 3,5-biphosphate (PI(3,5)P2) and phosphatidylinositol 4,5-biphosphate (PI(4,5)P2) in vitro (Kaminska et al., 2016). This domain is not related to known lipid-binding domains and is not sufficient to localize to PI(3)P-rich membranes in vivo. Notably, a causative chorea-acanthocytosis VPS13A point mutation (I2771R) in the APT1 domain alters its affinity for PI(3)P in vitro (Rzepnikowska et al., 2017). Modeling the homologous mutation in yeast (I2749R) similarly reduced PI(3)P binding. This mutant form of Vps13 was unable to rescue vacuolar protein sorting defects and drug sensitivities of vps13Δ null yeast suggesting that this domain is important for VPS13 protein function. The VPS13A APT1 interaction with PI(3)P-rich liposomes requires bivalent cations such as calcium or magnesium, while its interaction with PI(5)P-rich liposomes does not (Kolakowski et al., 2020). The cations are suggested to bind to the C-terminus of the APT1 domain as C-terminal truncations result in a loss of requirement for bivalent cations (Kolakowski et al., 2020). Interestingly, the VPS13A chorea-acanthocytosis mutation in the APT1 domain enhanced interactions with PI-rich liposomes in the presence of calcium ions. This altered specificity of the APT1 domain for certain phosphatidylinositol species could provide a method of regulating 22 recruitment to specific membranes in response to signaling events. Loss of the ability to respond to such signaling could result in VPS13 protein mislocalization and reduced lipid transport at MCSs. 1.6.4 Conserved amphipathic helices in ATG2_C domains VPS13 proteins have conserved autophagy-related protein C-terminal (ATG2_C) domains at their C-termini. These regions contain two to three predicted amphipathic helices (AHs), structures that contain hydrophobic residues on one helix face and polar residues on the other (Drin et al., 2007). This unique structure allows AHs to insert into membranes, sense regions of membrane curvature and even deform membranes (Giménez-Andrés et al., 2018). In human VPS13A and VPS13C, the ATG2_C region is sufficient for localization to lipid droplets (LDs) and, in VPS13A, to mitochondria as well (Kumar et al., 2018). Disruption of the hydrophobic face of the predicted amphipathic helix in a VPS13A ATG2_C domain-containing fragment blocked localization to LDs and mitochondria, suggesting the AHs are required for membrane recruitment. In line with this reasoning, recent in silico modeling has now proposed that the ATG2_C region forms a bundle of large AHs that can directly insert into the outer leaflets of organelles (Dall’Armellina et al., 2023). Here, the modeled membrane insertion was shown to disorder the lipid surface which may be important for lowering the energy requirements needed for lipid delivery or desorption at target membranes. 1.6.5 Pleckstrin homology domain The final C-terminal domain of VPS13 proteins is the Pleckstrin homology (PH) domain which was identified through structural homology searches (Söding et al., 2005; Fidler et al., 23 2016). Most PH domains interact with PIPs providing a mechanism for membrane targeting (Blomberg et al., 1999). A C-terminal Vps13 fragment containing both ATG2_C and PH domains binds PI(4,5)P2 in vitro (De et al., 2017; Kolakowski et al., 2021). Earlier studies had found that a dimer of the yeast Vps13 PH domain weakly localizes to yeast bud necks, and that expression of a mutant Vps13 protein lacking the PH domain was defective in NVJ localization in stationary phase yeast (Fidler et al., 2016) suggesting this domain does not bind strongly to lipids or membranes, but may contribute to membrane recruitment through coincidence detection. More recently, this domain was shown to interact with the GTPase Arf1 and to localize to the Golgi apparatus in a Arf1-dependent manner (Kolakowski et al., 2021). In human VPS13A, the PH domain has also been shown to form interactions with the scramblase XK and mutations that truncate this domain are causative for chorea-acanthocytosis (Dobson-Stone et al., 2002; Tomiyasu et al., 2011; Guillén-Samander et al., 2022). The role of the PH domain in other human VPS13 proteins has not been extensively studied, but they may similarly contribute to membrane targeting through interactions with protein partners. 1.7 Function of Vps13 at specific contact sites Although a number of localizations have been determined for both yeast and human VPS13 proteins, the full list of MCSs occupied by each homolog is not clear. The single organelle localizations listed in Table 1.1 and Table 1.2 are expected to be just one of two organelles tethered by VPS13 at MCSs. In this section, the possible organelle pairs tethered by VPS13 proteins will be discussed along with the evidence supporting a function in lipid transport at these sites. 24 1.7.1 Vps13 at the NVJ In yeast, Vps13 localizes to NVJs during glucose starvation (Lang et al., 2015), but its function at this contact site is unclear. The NVJ has been implicated in piecemeal microautophagy of the nucleus and lipid droplet biogenesis (Roberts et al., 2003; Kvam and Goldfarb, 2007; Hariri et al., 2018, 2019), but a functional role in these processes for Vps13 or its NVJ adaptor Ypt35 has not been uncovered. The nuclear ER and vacuole are a well-defined pair of organelles that are bridged by Vps13. Loss of the vacuolar adaptor, Ypt35, causes Vps13 to redistribute in the ER in agreement with the sufficiency of the Vps13 N-terminus for ER-targeting (Bean et al., 2018; Nakamura et al., 2021). Conversely, loss of Vps13 causes Ypt35 to become more evenly distributed around the vacuolar membrane. Thus, the interaction between Vps13 and Ypt35 clusters both proteins at the NVJ, but is not required for the association of these individual proteins with the ER or vacuole, respectively. 1.7.2 Vps13 at mitochondrial contact sites Vps13 is suggested to function in lipid transport at mitochondria based on the study of spontaneously-occurring suppressor mutations that allow Vps13 to compensate for loss of the ER-mitochondrial tether ERMES (Park et al., 2016; Lang et al., 2015). ERMES consists of 4 subunits, 3 of which have lipid-transporting SMP domains (Kornmann et al., 2009; Kopec et al., 2010; Kawano et al., 2018). Loss of ERMES subunits results in slow growth on fermentable media, abnormal, globular mitochondrial morphology and reduced lipid transport between ER and mitochondria (Kornmann et al., 2009). The slow growth and mitochondrial morphology defects caused by loss of two different ERMES subunits are rescued by dominant mutations in Vps13 but not by expression of a 25 synthetic ER-mitochondrial tether (Lang et al., 2015; Park et al., 2016; Kornmann et al., 2009), suggesting Vps13 does not simply hold organelles together but also restores impaired lipid transport. The basis for this suppression is unclear. One hypothesis is that the suppressor mutations alter Vps13 localization, and thus the loss of Vps13 from some contact sites provides a greater pool available to function elsewhere. Indeed, many of these mutations specifically block the localization of Vps13 to the NVJ in glucose limiting conditions (Park et al., 2016; Lang et al., 2015). An alternate explanation is that the mutations alter the lipid specificity of Vps13 as the dominant mutations largely affect residues that are predicted to reside at the center of the characteristic lipid transfer groove (Hanna et al., 2023). It is clear that Vps13 localization to mitochondrial contact sites is required for rescue (John Peter et al., 2017; Bean et al., 2018). ERMES suppression requires that mutant forms of Vps13 bind the mitochondrial adaptor, Mcp1 (John Peter et al., 2017; Bean et al., 2018). High levels of Mcp1, which redirect wild type Vps13 from endosomal puncta to the surface of mitochondria, also compensate for ERMES defects. Both the Vps13 suppressor mutations and overexpression of Mcp1 may thus increase the levels of Vps13 available to function at a mitochondrial contact site. Mitochondria make contacts with many other organelles including the ER, endosome, and vacuole (Lackner, 2019). The ability of Vps13 to rescue ERMES mutants was suggested to stem from a role in lipid transport at vacuole-mitochondria contacts (vCLAMPs), due to a requirement for the vCLAMP component Vps39 (Lang et al., 2015). In this model, lipids travel from ER to mitochondria by way of the vacuole, bypassing the ER-mitochondrial contact site. However, using separation-of-function alleles it was determined that the Vps39 function in vCLAMP formation is not important for this rescue. Vps39 is also a subunit of the homotypic 26 fusion and vacuole protein sorting (HOPS) tethering complex, and it is the role of Vps39 as a HOPS subunit that – together with the other HOPS subunits – is required for suppression of ERMES mutants by dominant Vps13 alleles (González Montoro et al., 2018). Vps13 is not observed at Vps39-containing vCLAMPs (González Montoro et al., 2018) but may form a different type of contact between mitochondria and vacuoles. Further work will be needed to understand how dominant suppressor mutations alter Vps13 localization or function. 1.7.3 Vps13 and prospore membrane expansion Vps13 localizes to the prospore membrane during sporulation where it is required for prospore membrane expansion and closure, and the formation of viable spores (Park and Neiman, 2012). Similar defects in sporulation are seen in mutants lacking Spo71, the Vps13 prospore membrane adaptor (Park et al., 2013). The dramatic expansion of the prospore membrane that encircles each prospore nucleus is reminiscent of the Atg2-mediated expansion of the phagophore membrane during autophagy, underlining the need for bulk lipid transport to drive this expansion. The small prospore membranes formed in vps13Δ mutants have reduced levels of PA, PI(4)P and PI(4,5)P2 suggesting alterations in lipid homeostasis (Park and Neiman, 2012). The opposing organelle at Vps13-prospore membrane contact sites is unknown. One suggested source of lipids for prospore membrane expansion is LDs (Hsu et al., 2017). LDs accumulate at contacts with growing prospore membranes and these docked LDs decrease in size as prospore membrane expansion proceeds. During meiosis II, levels of triacylglycerol (TAG) decrease while free fatty acids increase suggesting lipolysis occurs, and severe defects in spore viability are seen in cells lacking LDs (Hsu et al., 2017). It is thus interesting to hypothesize that 27 Vps13 tethers LDs to growing prospore membranes to provide a lipid source for membrane expansion. Because Vps13 is not known to transport free fatty acids, coordinated phospholipid synthesis coupled to the release of fatty acids from LDs could provide a source of transportable lipid at these sites. A similar mechanism drives autophagosome membrane formation, where activation of fatty acids from LDs and localized phospholipid synthesis is required for efficient phagophore elongation and autophagy (Schütter et al., 2020). Cells lacking LDs or with defects in lipases or phospholipid synthases exhibit autophagy defects, indicating release of fatty acids from LDs may be required for efficient autophagosome membrane expansion (Andrejeva et al., 2019; Dupont et al., 2014; Shpilka et al., 2015; Kannan et al., 2017). This suggests an interesting model, where enzymes involved in fatty acid activation and phospholipid synthesis are scaffolded with LDs at sites of membrane expansion to provide a pool of newly formed phospholipids for transport into growing autophagosome membranes. As Atg2 is the major lipid transport protein at these sites (Osawa et al., 2019; Valverde et al., 2019), it is interesting to speculate that Vps13 also scaffolds LDs and lipid biosynthetic enzymes at other contact sites to provide a source of glycerophospholipids for transport to growing membranes. 1.7.4 Vps13 at autophagosomal membranes Recently, yeast Vps13 was determined to function in cortical ER-phagy, but not in nuclear ER-phagy or bulk autophagy (Chen et al., 2020). In vps13Δ yeast the degradation of cortical ER markers is defective in cells treated with rapamycin, a potent inducer of autophagy. In this null mutant, ER membranes accumulate in late endosomes but are not packaged into autophagosomes or delivered to vacuoles (Chen et al., 2020). Upon treatment with rapamycin, 28 Vps13 was found to relocalize from endosomal puncta to vacuolar rims. Interestingly, this vacuolar localization occurred even in the absence of the endosomal/vacuolar adaptor, Ypt35 (Chen et al., 2020). It will be interesting to learn more about the role of Vps13 in ER-phagy, and if yeast Vps13 is involved in other organelle-specific autophagic processes. In other organisms, VPS13 homologs have an established role in autophagy. In the slime mold Dictyostelium discoideum, mutations in the VPS13 homolog TipC reduce autophagic flux (Muñoz-Braceras et al., 2015). In Drosophila melanogaster, mutation of the VPS13A/VPS13C homolog causes sensitivity to proteotoxic stress and an accumulation of protein aggregates that is rescued by overexpression of human VPS13A, suggesting a defect in delivery of autophagosomal membranes to lysosomes (Vonk et al., 2017). VPS13A has been shown to function in organelle-specific autophagy in human cell lines. VPS13A silencing results in impaired lysosomal degradation with accumulation of autophagic substrates (Muñoz-Braceras et al., 2019), mitochondrial fragmentation and reduced mitophagy (Yeshaw et al., 2019). Surprisingly, yeast Vps13 is not required for mitophagy and in fact deletion of VPS13 has been found to upregulate mitophagy in yeast (Chen et al., 2020; Park et al., 2016). As further evidence for a role of VPS13A in autophagy, erythrocytes from chorea-acanthocytosis patients accumulate autophagosomal cargo proteins. These patient erythrocytes have delayed mitochondrial and lysosomal clearance indicative of impaired autophagy (Lupo et al., 2016). Further work is required to determine if all VPS13 proteins are involved in autophagy, at what stage in autophagy they function, and which organelle contact sites are relevant for these functions. 29 1.7.5 VPS13 proteins at lipid droplets Although yeast Vps13 has not been found at LDs, there is abundant evidence linking human VPS13 proteins to LD function. Mammalian VPS13A and VPS13C localize to ER-LD contacts, with enhanced LD association seen for VPS13A in cells treated with oleic acid, an inducer of LD formation (Yeshaw et al., 2019; Kumar et al., 2018). VPS13A and VPS13C localize to ER-LD contacts through FFAT motifs that bind VAP at the ER, and AHs that target LDs (Kumar et al., 2018). The conserved AHs in Atg2 similarly localize to LDs (Tamura et al., 2017; Velikkakath et al., 2012), while its N-terminal chorein domain binds directly to ER membranes (Kotani et al., 2018a). The chorein and AH domains are conserved and it is plausible that the yeast Vps13 protein could similarly tether ER and LDs at either end of its lipid transport channel. Studies reporting changes in lipid droplet size, number, and motility in VPS13 and ATG2 mutants suggest they function at LDs. Loss of ATG2A and ATG2B increases the number and size of LDs (Velikkakath et al., 2012), while knockdown of VPS13A in mammalian cells, or of Vps13 in Drosophila, similarly increases LD number (Yeshaw et al., 2019). The reduced motility of VPS13A-associated LDs suggests VPS13A has a role in tethering LDs to another organelle. Notably, VPS13C is highly expressed in adipose tissue, particularly during adipocyte differentiation where VPS13C localizes to the surface of LDs (Yang et al., 2016; Ramseyer et al., 2018). Knockdown of VPS13C during adipocyte differentiation impairs adipogenesis (Yang et al., 2016), and VPS13C silencing in brown adipose tissue reduces LD size (Ramseyer et al., 2018). Whether all VPS13 proteins localize to ER-LD contacts and whether localization to these contact sites only occurs under certain conditions are important outstanding questions. 30 1.7.6 Vps13 at other contact sites It is likely that yeast Vps13 works at other contact sites that have not been identified. For example, Vps13 has a well-established role in the sorting pathway that transports CPY from the Golgi to the vacuole (Bankaitis et al., 1986). Vps13 has also been observed at peroxisomes (John Peter et al., 2017), but has not yet been implicated in lipid transport at this organelle. It is also possible that the role of Vps13 in some processes, including trans-Golgi network fusion, may not be related to contact sites (De et al., 2017). In other cases, Vps13 may be present at three-way junctions between organelles, making it difficult to ascertain which membranes are being tethered based on standard fluorescent microscopy. For example, Lam6, a sterol transport protein that is anchored to the ER by an N-terminal transmembrane domain, has been localized at ER-mitochondria contacts, vCLAMPs and NVJs (Elbaz-Alon et al., 2015). In such cases, the apparent localization to vCLAMPs could result from a 3-way junction that links ER-mitochondria and mitochondria-vacuole membranes. It is also possible that, due to its multiple membrane targeting domains, Vps13 can make contact with all three organelles at these junctions. Thus, much work is needed to determine which pairs of membranes are tethered by Vps13 at contact sites, and to identify the full set of contact sites where Vps13 functions. 1.8 Research objectives VPS13 proteins constitute a new class of lipid transfer proteins that simultaneously bridge organelle membranes at contact sites and channel bulks lipids between them. The four human VPS13 proteins localize to different subsets of MCSs and mutations in each gene are associated with distinct neurological disorders. One hypothesis that can explain the diverse 31 disorders resulting from the mutations in each VPS13 homolog is that loss of lipid transport at specific MCSs underlies this dysfunction. Therefore, understanding how VPS13 proteins target membranes and determining their sites of action are key to developing a better understanding of the associated pathologies. Using yeast Vps13 as a model for identifying membrane targeting determinants, our lab identified Ypt35 and Mcp1 as adaptor proteins that recruit Vps13 to endosomes/vacuoles and mitochondria respectively (Bean et al., 2018). Elaborating on this work in Chapter 2, I determine the necessity of a related motif in all three known adaptors for interaction with Vps13. Further I demonstrate that all adaptors interact at a previously uncharacterized six-repeat domain, which we have named the Vps13 Adaptor Binding (VAB) domain, in a motif-dependent manner. In Chapter 3, I detail experiments to dissect adaptor interactions with the VAB domain which led to the discovery of a single adaptor-binding pocket. Further, I demonstrate that modeling a spastic ataxia VPS13D missense mutation within this pocket blocks adaptor binding and Vps13 membrane recruitment in yeast, providing evidence that this domain has a conserved function in human homologs. After structural studies provided evidence for the novel extended lipid transfer channel of Vps13, we took advantage of the remote protein homology detection server, HHpred, to identify additional putative homologs with related functions. In Chapter 4, I describe my identification of Fmp27/Hob1 and its paralog Ypr117w/Hob2 as structurally related members of the Vps13/Atg2 family of bridging lipid transporters. I present our findings demonstrating that the N-terminally anchored Fmp27 localizes to ER-PM contacts through interactions with the uncharacterized protein Ybl086c, which we have named Hobbit interactor 1 (Hoi1). ER-PM recruitment of Fmp27 and Hoi1 is shown to be interdependent, and a large interaction interface is identified 32 through mutational analysis guided by AlphaFold2 protein structure prediction. Importantly, I demonstrate that loss of both paralogous Fmp27 and Hob2, or their PM interactor Hoi1, result in enhanced sensitivity to sterol-binding drugs and a redistribution of accessible sterols from the cytosolic PM leaflet to intracellular puncta providing support that these putative lipid transfer channels affect lipid homeostasis and distribution. 33 Chapter 2: Competitive organelle-specific adaptors recruit Vps13 to membrane contact sites 2.1 Prelude The work described in this chapter is a portion of a larger collaborative manuscript published as: Bean, B.D.M., S.K. Dziurdzik, K.L. Kolehmainen, C.M.S. Fowler, W.K. Kwong, L.I. Grad, M. Davey, C. Schluter, and E. Conibear. 2018. Competitive organelle-specific adaptors recruit Vps13 to membrane contact sites. Journal of Cell Biology. 217:3593–3607. doi:10.1083/jcb.201804111. Additionally, the final figure in this section is modified from: Dziurdzik, S.K., B.D.M. Bean, and E. Conibear. 2018. An interorganellar bidding war: Vps13 localization by competitive organelle-specific adaptors. Contact. 1. doi:10.1177/2515256418814621. In this chapter, I present my work establishing the necessity of a shared motif in adaptor proteins for interaction with Vps13 and identify a shared binding site for all adaptors in a conserved six-repeat region of Vps13. For brevity and focus, I summarize relevant findings of other lab members in identifying the Vps13 endosomal and vacuolar adaptor, demonstrating that adaptors are sufficient for Vps13 recruitment, and in establishing that adaptors directly compete for Vps13 recruitment. Additional information on this work can be accessed in the published journal article. Of note, at the time the work described here was performed, little was known about the function or structure of Vps13. Due to this, the chapter is written in the perspective of the time of the study. At that time, the human VPS13 proteins had not yet been observed at membrane contact sites, nor were the VPS13 membrane binding domains described in Chapter 1 34 demonstrated to localize to membranes. Additionally, the chorein domain of yeast Vps13 had not yet been structurally resolved and there was little evidence that VPS13 proteins could directly transport lipids between membranes. 2.2 Introduction Membrane contact sites (MCSs), regions where two organelles come within close proximity without fusing, are critical regions of lipid and ion transport, lipid biosynthesis and interorganellar communication (Eisenberg-Bord et al., 2016; Phillips and Voeltz, 2016; Helle et al., 2013). These sites are formed by protein tethers that span the 10-30 nm distance between opposing membranes and link them through transmembrane anchors, lipid-binding domains and/or protein interactions (Eisenberg-Bord et al., 2016; Gatta and Levine, 2017). The extent of any given contact site is modulated in response to cellular cues and changing metabolic needs, and the relative size of different membrane contacts is tightly coordinated (Elbaz-Alon et al., 2015, 2014; Liu et al., 2016; Bian et al., 2018; Murley and Nunnari, 2016). The expansion or contraction of contact sites requires dynamic changes in the localization of tethers and lipid transfer proteins, but how these changes are coordinated is not well understood. In S. cerevisiae, protein tethers have been identified that create membrane contact sites connecting the ER, mitochondria and vacuole. The endoplasmic reticulum-mitochondria encounter structure (ERMES) complex tethers the ER to mitochondria (Kornmann et al., 2009), Vps39 drives formation of the vacuole-mitochondria patch (vCLAMP) (Hönscher et al., 2014; Elbaz-Alon et al., 2014), and Nvj1 and Vac8 link the ER to the vacuole to form the nucleus-vacuole junction (NVJ) (Pan et al., 2000). These contact sites may functionally overlap as loss of mitochondrial contact sites caused by defects in the ERMES complex causes expansion of 35 mitochondrial-vacuole (vCLAMP) contacts, which may provide an alternative pathway for transporting lipids to mitochondria (Elbaz-Alon et al., 2014; Hönscher et al., 2014). Identifying the constituent proteins that regulate contact site assembly and function and understanding how their localization is controlled remains a major challenge in the field. Vps13 is a highly conserved 358 kDa protein that is present at multiple cellular locations including vCLAMP and NVJ MCSs, endosomes, and the prospore membrane (Huh et al., 2003; Lang et al., 2015; Park et al., 2013, 2016). Although Vps13 in yeast and other organisms has been implicated in processes including vesicle fusion, autophagy and actin regulation (De et al., 2017; Brickner and Fuller, 1997; Dalton et al., 2017; Muñoz-Braceras et al., 2015; Föller et al., 2012; Rzepnikowska et al., 2017), its roles at membranes, including membrane contact sites, are not known. Humans contain four VPS13 proteins (VPS13A-D) that when non-functional cause different diseases - chorea-acanthocytosis, Cohen syndrome, Parkinson’s disease and spastic ataxia, respectively - suggesting they have functionally diverged (Rampoldi et al., 2001; Kolehmainen et al., 2003; Majounie et al., 2016; Seong et al., 2018; Drin et al., 2007). Although mammalian VPS13 proteins have not yet been found at contact sites, dysfunction of membrane contact sites is implicated in neurological and movement disorders (Paillusson et al., 2016; van der Kant and Neefjes, 2014). The location of yeast Vps13 at contact sites is regulated by nutrient availability. During mitotic growth when glucose is abundant, Vps13 is found primarily at endosomes and is frequently observed near endosome-mitochondria contacts, but relocalizes to the NVJ when glucose is limiting (Lang et al., 2015; Park et al., 2016). Vps13 is also reported to localize to vCLAMPs and this is thought to be essential for cell viability when ERMES is disrupted (Lang et al., 2015). Spontaneously occurring dominant mutations in Vps13 suppress ERMES defects, 36 and may favor relocalization to vCLAMPs (Lang et al., 2015; Park et al., 2016). This discovery highlights the critical role of Vps13 at membrane contact sites and suggests that uncovering the mechanisms that control Vps13 localization may be key to understanding coordination between contact sites. Vps13 binds phosphatidic acid and phosphoinositides in vitro (De et al., 2017; Rzepnikowska et al., 2017), but it is unclear if lipid binding domains are sufficient for targeting to membranes in vivo. Instead, Vps13 may rely on interactions with membrane-specific adaptor proteins for recruitment. At the time the research described in this chapter was performed, the only known adaptor was the meiosis-specific protein Spo71, which recruits Vps13 to the prospore membrane, where it facilitates prospore membrane expansion and closure (Park et al., 2013; Park and Neiman, 2012). We had hypothesized that if one adaptor can regulate Vps13 localization, additional adaptors may exist for recruiting Vps13 to other organelle membranes. Indeed, in our publication the work of our lab identified the uncharacterized sorting nexin, Ypt35, as a new Vps13 adaptor protein (Bean et al., 2018). We demonstrate that Ypt35 is required for Vps13 recruitment to endosomes and, under glucose-limiting conditions, to the vacuole membrane at NVJ contact sites, and determine that this recruitment requires a conserved N-terminal motif in Ypt35 (Bean et al., 2018). In this chapter, we map the Ypt35 binding site on Vps13 to a previously uncharacterized highly conserved 6-repeat domain. Interaction with this domain is dependent on the conserved motif in Ypt35. By searching for other proteins that contain similar sequences, we identify related motifs in the meiosis-specific adaptor Spo71 and the mitochondrial membrane protein Mcp1. We determine that the related motifs of all 3 adaptors are necessary for interaction with 37 Vps13, and find they interact with the same region suggesting these adaptors compete for binding to regulate the localization of Vps13 to different organelle membranes. 2.3 Results 2.3.1 The Vps13 DUF1162 domain binds the Ypt35 PxP motif In our publication, we had identified the PI(3)P-binding Phox homology (PX) domain-containing protein Ypt35 as the endosomal and vacuolar Vps13 adaptor (Bean et al., 2018). We mapped the Ypt35 site necessary for Vps13 recruitment to a conserved region in the unstructured N-terminus between residues 5 and 15 which has a consensus sequence of ΦxxΦxPxPΦxΦ, where Φ is a hydrophobic amino acid, in Saccharomycetales-level homologs. Mutations of individual conserved residues within this motif disrupted Vps13 recruitment, however substitution of both invariant prolines to alanines (P10A, P12A) abolished recruitment to background levels (Bean et al., 2018). Due to the necessity of the prolines for Vps13 recruitment, we refer to the interaction motif as the “PxP” motif. Vps13 contains several conserved domains which could mediate interactions with the Ypt35 PxP motif. These include an N-terminal chorein domain, a central domain of unknown function (DUF1162), an APT1 domain, two ATG2 C-terminal domains and a putative lipid-binding Pleckstrin homology (PH) domain (Velayos-Baeza et al., 2004; Rzepnikowska et al., 2017; Fidler et al., 2016). To identify the part of Vps13 that interacts with Ypt35, we created a series of Vps13-GFP (green fluorescent protein) truncations that separate these Vps13 domains (Fig 2.1A). To detect the interaction in the absence of crosslinkers, the Vps13 fragments and Ypt35 were overexpressed from the ADH1 promoter. All truncated proteins were expressed, but only truncations containing the DUF1162 domain or to a lesser extent the C-terminus were able 38 Figure 2.1: A central region of Vps13 binds Ypt35 in a PxP motif-dependent manner. (A) Schematic of Vps13 truncations with conserved domains labelled. (B) Co-immunoprecipitation of WT or mutant Ypt35-3HA with Vps13-GFP fragments indicates that interactions with DUF1162-containing regions are proline dependent. IP, immunoprecipitation; WCL, whole-cell lysate. (C) Densitometry of C; n = 3. Error bars indicate SEM. (D) Ypt35-3HA binds the minimal DUF1162-containing Vps13(1732–2556)-GFP fragment. ADH1pr-driven WT or truncated Vps13-GFP was immunopurified, and coprecipitating ADH1pr-expressed Ypt35-3HA was detected by immunoblot. (E) Schematic of S. cerevisiae Vps13, highlighting an extended DUF1162 domain defined by six repeats identified using HHrepID (Biegert and Söding, 2008). (F) An alignment of the 39 most conserved section of the S. cerevisiae DUF1162 repeats. The highly conserved invariant asparagine of each repeat is denoted with an arrow. to co-purify Ypt35-3HA (3xHemagglutinin) (Bean et al., 2018, Fig 2.1B,C). While the interaction with the C-terminus was reproducible, it was relatively weak and a fragment containing both the DUF1162 domain and the C-terminus did not display enhanced binding. In contrast, interactions with DUF1162-containing fragments were more robust. A smaller DUF1162-containing fragment, Vps13(1732-2556), was unstable but able to co-purify Ypt35-3HA (Fig 2.1D), suggesting the PxP motif binds Vps13, directly or indirectly, at or near the DUF1162 domain. To test the importance of the PxP motif for these interactions, we tested binding of a subset of the Vps13 fragments to the Ypt35(P10,12A)-3HA mutant (Fig 2.1B,C). Full-length Vps13-GFP, or DUF1162-containing Vps13-GFP truncations, were dependent on the conserved prolines for interactions with Ypt35. In contrast, binding to the C-terminal fragment was largely PxP-independent. These results strongly suggest that the Ypt35 PxP motif binds Vps13 either at the DUF1162 domain or at an adjacent region within the smallest interacting DUF1162-containing fragment. 2.3.2 The DUF1162 domain is part of a 6-repeat region Ypt35 is the second DUF1162-interacting protein to be identified. Previously, yeast two-hybrid and split yellow fluorescent protein (YFP) experiments in A. thaliana indicated that the Vps13 homolog SHRUBBY interacts with the SHORT-ROOT (SHR) transcription factor through the DUF1162 domain, now commonly referred to as a SHR binding domain (SHR_BD) (Koizumi and Gallagher, 2013). Analysis of the smallest Ypt35-interacting Vps13 fragment 40 (1732-2556) using HHrepID (Biegert and Söding, 2008) identified six high probability repeats spanning residues 1852-2522, roughly twice the length of the classic DUF1162 domain (2206-2484) (Fig 2.1E). The first ~25 residues of each ~100 amino acid repeat were the most highly conserved and contained an invariant asparagine residue (Fig 2.1F). The DUF1162 repeats are also present in human VPS13A/B/C/D genes. Similar to yeast Vps13, the N-terminal region of each repeat is the most conserved and contains a nearly invariant asparagine residue (Bean et al., 2018). All human VPS13 genes encode 6 repeats, but two VPS13B repeats do not contain the highly conserved asparagine residue. We suggest that this extended repeat structure represents a complete DUF1162 domain. 2.3.3 Spo71 binds the Vps13 DUF1162 domain through a PxP motif Having determined that Ypt35 interacts with the DUF1162 domain-containing region of Vps13 we wondered if Spo71, an established Vps13 prospore membrane adaptor, binds Vps13 in a similar manner. Using the same panel of overexpressed Vps13-GFP truncations (Fig 2.1A), we found that Spo71 also binds DUF1162 domain-containing fragments and, to a lesser extent, the C-terminal domain (Fig 2.2A,B). To identify candidate PxP-based binding motifs in Spo71, we generated a Multiple Expectation Maximization Algorithms for Motif Elucidation (MEME) motif (Bailey and Elkan, 1994) from Saccharomycetales-level Ypt35 homologs and used it in a Find Individual Motif Occurrences (FIMO) (Grant et al., 2017) search of the S. cerevisiae proteome. Among the hits, only one was identified within Spo71 at residues 385-399 (Fig 2.2C). This hit matched the Ypt35 PxP motif consensus with the exception of a threonine in one hydrophobic position. This motif was determined to be sufficient for Vps13 recruitment by using a chimeric recruitment assay where the putative Spo71 PxP motif was fused to a red fluorescent 41 Figure 2.2: The Vps13 DUF1162 domain recognizes the meiosis-specific adaptor Spo71 through a PxP motif. (A) Co-immunoprecipitation of TEF1pr-driven Spo71-3HA with ADH1pr-driven Vps13-GFP fragments indicates that fragments containing the DUF1162 domain are sufficient for interaction. (B) Densitometry of selected strains from A; n = 3. Error bars indicate SEM. (C) A schematic of Spo71 highlighting a putative Vps13 interaction motif identified by a FIMO search using a MEME motif generated from Ypt35 homologues. The threonine residue in the Spo71 sequence that does not correspond to the Ypt35 homolog consensus is underlined in red. (D) ADH1pr-driven Vps13-GFP was immunopurified and coprecipitating WT or mutant TEF1pr-expressed Spo71-3HA was detected. IP, immunoprecipitation; WCL, whole-cell lysate. (E) Densitometry of D; n = 3. Error bars indicate SEM. 42 protein (RFP)-labelled PI(3)P-binding FYVE domain of EEA1 and assessing its ability to recruit Vps13 to endosomal puncta in ypt35Δ cells (Bean et al., 2018). The presence of a polar threonine at a conserved position in the Vps13 interaction motif indicates there is greater amino acid variability than our initial analysis suggested. Although a chimera containing the Spo71 PxP motif recruited Vps13 to membranes, full-length Spo71 may contain redundant signals. We used co-immunoprecipitation to determine if the motif at residues 385-399 is necessary for Vps13-Spo71 interaction. While wild type Spo71-3HA efficiently co-purified with Vps13-GFP, the interaction was severely disrupted by mutation of the invariant prolines (P390A, P392A), and abolished by deletion of the full motif (Fig 2.2D,E). A low level of binding to the Spo71(P390,392A) mutant indicates that other residues, such as the conserved hydrophobic amino acids, may be capable of residual binding in the absence of the invariant prolines. Our results suggest Spo71 contains a PxP motif that is necessary and sufficient for Vps13 binding, indicating that this mechanism for Vps13 interaction is conserved between proteins. 2.3.4 Mcp1 is a mitochondrial PxP motif-containing Vps13 adaptor We reasoned that a search of the yeast proteome for sequences similar to the PxP motif might identify additional Vps13 adaptors. We pooled Saccharomycetales-level motif alignments from Ypt35 and Spo71 to generate a new MEME motif (Fig 2.3A). A FIMO search of the S. cerevisiae proteome using this motif identified 296 hits at a p<1x10-4 threshold (Bean et al., 2018), which included a putative PxP motif in the outer mitochondrial membrane protein Mcp1 (p<3.7x10-5). Mcp1 was a strong adaptor candidate as dominant Vps13 alleles suppress 43 Figure 2.3: Mcp1 is a mitochondrial PxP motif-containing Vps13 adaptor. (A) Schematic of Mcp1 highlighting a putative PxP interaction motif identified by FIMO using a MEME motif generated from Ypt35 and Spo71 homologues. (B) The motif is required for co-immunoprecipitation of Vps13 and Mcp1. Vps13-GFP and WT and mutant forms of Mcp1-3HA were overexpressed from the ADH1 promoter. IP, immunoprecipitation; WCL, whole-cell lysate. (C) Densitometry of B; n = 3. Error bars indicate SEM. (D) Mcp1-3HA interacts with DUF1162–containing Vps13 fragments but not a C-terminal fragment in a PxP motif–dependent manner. ADH1pr-driven Vps13-GFP was immunopurified, and coprecipitating WT or mutant ADH1pr-overexpressed Mcp1-3HA was detected. IP, immunoprecipitation; WCL, whole-cell lysate. ERMES mutants only when Mcp1 is present, and high levels of Mcp1 also suppress ERMES defects (Lang et al., 2015; Tan et al., 2013). 44 The predicted PxP motif in Mcp1, 3KLHEVPPEPV12, is conserved in other fungal species and resides in a predicted 58 residue N-terminal cytosolic domain that is followed by five predicted transmembrane helices (Fig 2.3A). Using a series of Mcp1-RFP-FYVE chimeras we determined this motif was both necessary and sufficient for Vps13 recruitment (Bean et al., 2018). We next tested if we could observe a physical interaction between Vps13 and full-length Mcp1. Mcp1-3HA co-purified with Vps13-GFP, and this interaction was disrupted when either the putative motif was deleted or the core prolines were mutated to alanines (Fig 2.3B,C). Mcp1-3HA showed weak PxP-dependent binding to the Vps13(1363-2556)-GFP DUF1162 domain-containing fragment, while the low level of binding to the Vps13 C-terminal fragment was largely unaffected by mutation of the PxP motif (Fig 2.3D). Our results indicate that Mcp1 contains a bona fide PxP motif that is necessary and sufficient for interaction with Vps13. In our publication, we further demonstrated that Mcp1 functions as the Vps13 mitochondrial adaptor. High levels of Mcp1, but not a mutant form lacking the PxP motif, were sufficient to recruit Vps13 to mitochondria (Bean et al., 2018). Additionally, Mcp1 PxP-mediated Vps13 recruitment to mitochondria was confirmed to be necessary for rescue of mitochondrial morphology defects in mdm10Δ mutants lacking a subunit of the ERMES complex (Bean et al., 2018). This supports a shared role for Vps13 and Mcp1 in a potential lipid bypass pathway that can suppress mitochondrial defects in ERMES mutants. Altogether, these results show that Mcp1 recruits Vps13 to mitochondria through a related PxP motif for a putative role in lipid transfer. 45 2.3.5 Vps13 adaptors compete to recruit Vps13 to different membranes The observation that all three identified adaptors rely on related motifs for Vps13 recruitment and that these motifs interact with the same region of Vps13 suggested that the adaptors may compete for a shared binding site. Consistent with this hypothesis, we found that strong overexpression of soluble forms of Ypt35, Spo71 and Mcp1 PxP motifs fused to an RFP marker were sufficient to disrupt recruitment of Vps13 to endosomes and mitochondria in strains overexpressing Ypt35 or Mcp1 respectively (Bean et al., 2018). Automated quantitation of the relative Vps13 recruitment levels suggested that the Spo71 PxP motif had the strongest ability to disrupt recruitment, while the Mcp1 PxP motif had the least (Bean et al., 2018). Taken together, these results strongly suggest that Vps13 adaptors compete for the same binding site on Vps13 and further show that different adaptor PxP motifs have separate affinities for interaction, allowing changes in adaptor expression, or PxP motif exposure, to act as a mechanism for regulating Vps13 localization. 2.4 Discussion We have identified Ypt35 as the adaptor protein that recruits Vps13 to endosomal and vacuolar membranes and Mcp1 as the adaptor for Vps13 at mitochondria. Together with the prospore membrane adaptor Spo71, these proteins direct Vps13 to distinct cellular locations (Fig 2.4A). Remarkably, all three adaptors use a related motif to bind a conserved 6-repeat region that includes the DUF1162 domain. We recommend this region be renamed the VPS13 Adaptor Binding (VAB) domain. Our results suggest that Vps13 localization may largely be determined by competition between adaptors and thus changes in the level of a particular adaptor, or in the exposure of its recruitment motif, could drive the localization of Vps13 to MCSs in response to 46 environmental stimuli or altered lipid homeostasis. 2.4.1 PxP motif-containing adaptors compete for Vps13 binding We identified a short motif within the unstructured N-terminus of all three Vps13 adaptors that binds directly or indirectly to a conserved region of Vps13 containing the DUF1162 domain. The core motif present in all three adaptors consists of prolines flanked by hydrophobic residues that conform to a ΦxxΦxPxP(Φ/T) consensus (Fig 2.4B). In each adaptor, the motif is located in an unstructured region ~50-100 amino acids N-terminal to lipid binding or transmembrane domains, which may help position Vps13 relative to the membrane. The similarity between recruitment motifs suggests that adaptors compete for binding to a single site on Vps13. Indeed, we found that soluble fusion proteins containing these motifs can displace Vps13 from endosomes or mitochondria. Our competition experiments suggest Spo71 has the strongest affinity for Vps13, while Mcp1 has the weakest. Slight differences between motifs, such as the presence of hydrophobic or acidic residues following the PxP sequence, may explain these different signal strengths. The relatively high affinity of Spo71 for Vps13 could explain how Vps13 is efficiently relocalized to the prospore membrane during meiosis, when Spo71 expression is induced (Park et al., 2013). At steady state, the relative affinities of Mcp1 and Ypt35 motifs may dictate Vps13 distribution between endosomes and mitochondria. Furthermore, this distribution could be fine-tuned by changes in the relative abundance of Ypt35 and Mcp1, or posttranslational modifications that modulate the conformation or affinity of their binding motifs. Are there other adaptors? Vps13 can localize to peroxisomes (John Peter et al., 2017) and, although we found little evidence of Golgi targeting in nutrient-rich conditions (Bean et al., 47 Figure 2.4: Model of Vps13 recruitment by adaptor proteins with PxP motifs. (A) Ypt35, Spo71, and Mcp1 all contain PxP motifs that compete to recruit Vps13 from a cytosolic pool to endosomes/vacuole, the prospore membrane, and mitochondria, respectively. The PxP motifs bind to the six-repeat VAB domain (dark green lines) which extends beyond the previously described DUF1162 domain. Yeast Vps13 may interact with the ER directly or through an additional unidentified adaptor, allowing bridging of membranes at contact site that can aid in a putative lipid transfer function (gray arrows). The presence of Vps13 at contact sites that link 48 mitochondria or endosomes with the ER (dotted lines) is speculative. (B) WebLogo of the PxP interaction motif from Saccharomycetales-level homologs of Ypt35, Spo71, and Mcp1. The consensus is shown with Φ indicating hydrophobic residues. 2018), Vps13 is required for homotypic fusion of Golgi membranes and for Golgi/endosomal trafficking (De et al., 2017; Dalton et al., 2017; Brickner and Fuller, 1997). These functions may not involve MCSs (De et al., 2017; Dalton et al., 2017), but likely require association of Vps13 with membranes. A search of the yeast proteome successfully identified recruitment signals in Mcp1 and Spo71, suggesting this approach may identify new adaptors that use a PxP motif to recruit Vps13 to other membranes. 2.4.2 Possible bridging role for Vps13 at contact sites It remains to be determined if Vps13 acts as a tether, a scaffold, and/or a lipid-transfer protein. Vps13 does not act as a primary tether at NVJs as loss of Vps13 does not block NVJ formation (Bean et al., 2018). Vps13 may have a redundant tethering role at the NVJ and could also contribute to the ER-vacuolar non-NVJ sites that persist in nvj1 mutants (Murley et al., 2015; Henne et al., 2015; Wong and Levine, 2016). Our work demonstrated that Vps13 is necessary to enrich Ypt35 at NVJ contact sites (Bean et al., 2018) suggesting Vps13, or an additional interactor, can associate with the ER membrane thus bridging organelles at this MCS. Alternatively, Vps13 may not need to bridge two membranes and instead it could provide a scaffold to enrich lipid transfer proteins at membrane contact sites. Vps13 contains several domains that bind lipids (De et al., 2017; Rzepnikowska et al., 2017). While no direct role in lipid transfer has been demonstrated, it is interesting that John Peter et al. find a function for 49 Mcp1 in ERMES suppression beyond Vps13 recruitment and speculate that Vps13 shuttles lipids to Mcp1 (John Peter et al., 2017). Conserved lipid binding domains in Vps13 could work together with PxP sequences for membrane recruitment. The APT1 domain may bind PI(3)P to enhance endosomal targeting, while the C-terminal PH domain (Fidler et al., 2016) may contribute to Vps13 localization at mitochondria. Because all three adaptors displayed a weak PxP-independent binding to the Vps13 C-terminus, these domains may make other contacts with adaptors, or recruit cofactors. For example, the C-terminus of VPS13B binds Rab6 for recruitment to the Golgi in human cells (Seifert et al., 2015), suggesting combinatorial interactions with Rab GTPases or lipids may contribute to Vps13 localization. 2.4.3 The adaptor-binding domain of Vps13 is a conserved 6-repeat region We found the Vps13 Adaptor-Binding (VAB) domain is composed of 6 repeats spanning a region roughly twice the size of the previously defined DUF1162/SHR-BD domain. All 6 repeats, each featuring an invariant asparagine residue, are present in most VPS13 family members including human VPS13A, VPS13C and VPS13D. Although VPS13B contains 6 identifiable repeats, two lack the invariant asparagine. Mutation of the key asparagine in the most conserved repeat (N2993S) of VPS13B causes Cohen syndrome (Kolehmainen et al., 2003), while mutation of the invariant asparagine in the last repeat of VPS13D (N3521S) is associated with a childhood-onset movement disorder (Gauthier et al., 2018; Seong et al., 2018). Additionally, a VPS13A homozygous missense mutation of another highly conserved residue within the last repeat (W2460R) results in chorea-acanthocytosis and has been found to disrupt VPS13A localization to uncharacterized vesicular structures (Dobson-Stone et al., 2002; 50 Velayos-Baeza et al., 2008). This suggests the VAB domain repeats have a critical functional role, though candidate binding partners have yet to be identified. Human VPS13 proteins are ubiquitously expressed and localize to a variety of organelles including Golgi, lipid droplets, and mitochondria (Seifert et al., 2015; Yang et al., 2016; Ramseyer et al., 2018). Although they have not yet been demonstrated to be at contact sites, a recent study found VPS13C on a subdomain of lipid droplets proximal to mitochondria (Ramseyer et al., 2017). Moreover, VPS13A/C contain predicted FFAT motifs, suggesting they interact with VAMP-associated protein (VAP) proteins at the ER (Mikitova and Levine, 2012; Huttlin et al., 2015). The functional divergence of human VPS13 family members may allow them to fulfill specialized roles at different organelles involving unique adaptor repertoires, explaining how dysfunction of these related proteins can cause diverse diseases. Further studies in yeast promise to shed more light on this family of proteins. 2.5 Materials and methods 2.5.1 Yeast strains and plasmids Strains and plasmids are listed in Tables 2.1 and Table 2.2 respectively. Yeast strains were made by standard PCR-based homologous recombination (Janke et al., 2004; Longtine et al., 1998). Gene deletions were confirmed by colony PCR. Plasmids were made via homologous recombination in yeast by cotransforming linearized plasmids with PCR products. The plasmids were recovered in E. coli and sequenced. 51 Table 2.1: Yeast strains used in Chapter 2 Strain ID Genotype Source BY4742 BY4742 MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 Dr. E.K. O’Shea (Harvard University) KK16 BY4742 MATα kanMX4::ADH1-VPS13 his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK17 BY4742 MATα kanMX4::ADH1-VPS13-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) MDY1141 BY4742 MATα kanMX4::ADH1-VPS13(1-440)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) MDY1142 BY4742 MATα kanMX4::ADH1-VPS13(1-853)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK48 BY4742 MATα kanMX4::ADH1-VPS13(1-1362)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK19 BY4742 MATα kanMX4::ADH1-VPS13(1-2556)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK42 BY4742 MATα kanMX4::ADH1-VPS13(1363-2556)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK43 BY4742 MATα kanMX4::ADH1-VPS13(1732-2556)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK44 BY4742 MATα kanMX4::ADH1-VPS13(1363-3144)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) KK23 BY4742 MATα kanMX4::ADH1-VPS13(2557-3144)-GFP+::HIS3 leu2Δ0 lys2Δ0 ura3Δ0 (Bean et al., 2018) Table 2.2: Plasmids used in Chapter 2 Plasmid ID Plasmid Description Source pMD232 pRS415-ADH1-YPT35-3HA (LEU2, CEN) (Bean et al., 2018) pMD291 pRS415-ADH1-YPT35(P10,12A)-3HA (LEU2, CEN) (Bean et al., 2018) pMD166 pRS425-TEF1-SPO71-3HA (LEU2, 2μ) (Bean et al., 2018) pBB59 pRS425-TEF1-SPO71(P390,392A)-HA (LEU2, 2μ) (Bean et al., 2018) pBB60 pRS425-TEF1-SPO71(384-395Δ)-HA (LEU2, 2μ) (Bean et al., 2018) pBB44 pRS416-ADH1-MCP1-3HA (URA3, CEN) (Bean et al., 2018) 52 Plasmid ID Plasmid Description Source pBB56 pRS416-ADH1-MCP1(P9,11A)-3HA (URA3, CEN) (Bean et al., 2018) pBB45 pRS416-ADH1-MCP1(4-12Δ)-3HA (URA3, CEN) (Bean et al., 2018) 2.5.2 Co-immunoprecipitation Yeast were grown to log-phase in minimal synthetic dextrose media, 40 OD600 of yeast were harvested and spheroplasts prepared by incubating with zymolase (MJS BioLynx) at 30 °C. Spheroplasts were lysed in lysis buffer (50 mM HEPES, 0.1% Tween-20, 50 mM NaCl, 1mM EDTA, 100 mM PMSF, 1x HALT protease inhibitor cocktail; pH 7.4). The lysates were incubated with rabbit anti-GFP (Eusera, catalog # EU2) followed by protein A–Sepharose beads (GE Healthcare) at 4 °C. Washed beads were heated in sample buffer at 70 °C for 5 min. Samples were run on SDS-PAGE gels, transferred to nitrocellulose membranes and blotted with mouse anti-GFP (Roche, catalog # 11-814-460-001) or mouse anti-HA (Covance, catalog # MMS-101R) followed by goat anti-mouse conjugated to horseradish peroxidase (HRP; Jackson ImmunoResearch, catalog # 115-035-146) antibodies. West Pico (Pierce) chemiluminescent reagent was added to the blot and used to expose Amersham Hyperfilm (GE Healthcare). Films were scanned and densitometry was performed in Fiji (Schindelin et al., 2012). 2.5.3 MEME and FIMO-based motif search Saccharomycetales-level homologs of Ypt35 and Spo71 were identified by OrthoDB (Zdobnov et al., 2017) and aligned with MUSCLE (Edgar, 2004). Sequences that contained 3 or more residues consistent with the Ypt35/Spo71 consensus and that were not identical to either Ypt35 or Spo71 were selected. A region of 15-29 residues within the alignment centered on the core prolines was analyzed using Multiple Expectation Maximization Algorithms for Motif 53 Elucidation (MEME) (Bailey and Elkan, 1994). In each case we selected the most significant MEME motif as an input for a Find Individual Motif Occurrences (FIMO) search of the S. cerevisiae proteome with a threshold of P < 0.0001 (Grant et al., 2017). 54 Chapter 3: A VPS13D spastic ataxia mutation disrupts the conserved adaptor-binding site in yeast Vps13 3.1 Prelude The majority of the work described in this chapter has been published as: Dziurdzik, S.K., B.D.M. Bean, M. Davey, and E. Conibear. 2020. A VPS13D spastic ataxia mutation disrupts the conserved adaptor-binding site in yeast Vps13. Human Molecular Genetics. 29:635–648. doi:10.1093/hmg/ddz318. At the time of publication we had generated predicted models of VPS13 VAB domains using I-TASSER (Yang et al., 2015) which suggested they form a WD40-like fold, in agreement with other publications at the time (Kumar et al., 2018; Ugur et al., 2020). Our experimental findings were discussed in the context of these models in the published manuscript. However, after the release of the protein structure prediction algorithm, AlphaFold (Jumper et al., 2021), we were able to generate new models of the VAB domain with higher confidence that proposed a different structure. Notably, the thermophilic yeast Vps13 VAB domain and the adaptor binding site have now been structurally resolved by crystallography (Adlakha et al., 2022) and confirm these AlphaFold predictions. Our new AlphaFold models are presented in a separate manuscript under revision: Jeffers, K.R., Dziurdzik, S.K., M. Davey, and E. Conibear. 2023. Identification of a shared binding site for Vps13 organelle-specific adaptors. Molecular Biology of the Cell. In review. In this thesis chapter, the results and discussion of the original publication have been modified to consider only the AlphaFold models of the VAB domain. The new model is presented in Results subsection 3.3.7 and Figure 3.7. Discussion subsection 3.4.1 has been 55 modified to discuss the experimental findings in the context of the AlphaFold model and Figure 3.8 altered to match our new model of Vps13 membrane recruitment. The difference between structural models, as well as a comparison to the later published crystal structure, are discussed in more detail in Chapter 5. 3.2 Introduction Membrane contact sites (MCSs) — regions where organelle membranes are tethered together in close proximity— are key sites of non-vesicular lipid and ion transport (Scorrano et al., 2019; Wu et al., 2018). Dysfunction at these sites can perturb cellular functions including Ca2+ signaling, lipid transport, autophagy, intracellular protein transport and ER homeostasis (Paillusson et al., 2016; Henne, 2017; Krols et al., 2016). Indeed, a growing number of mutations in MCS proteins have been found to cause disease (Krols et al., 2016; Paillusson et al., 2016). A key example of disease-associated MCS proteins are the VPS13 paralogs, which were recently implicated in lipid transport at MCSs (Kumar et al., 2018). Autosomal recessive mutations in the four human VPS13 proteins (VPS13A-D) are associated with distinct neurological disorders. Mutations in VPS13A cause chorea-acanthocytosis (OMIM 200150), a progressive movement disorder with abnormal red blood cell morphology (Rampoldi et al., 2001). Mutations in VPS13B result in Cohen syndrome (OMIM 216550), a congenital multisystem disorder characterized by intellectual disability and developmental delay, and are implicated in autism (Kolehmainen et al., 2003; Yu et al., 2013; Balicza et al., 2019). More recently mutations have been identified in VPS13C and VPS13D that respectively result in rapidly progressive, early-onset Parkinson’s disease (OMIM 616840) (Lesage et al., 2016), and spastic ataxia (OMIM 607317) (Seong et al., 2018; Gauthier et al., 2018). Disease-causing 56 mutations in VPS13A-C are frequently null or truncating, resulting in no detectable protein (Dobson-Stone et al., 2002; Balikova et al., 2009; Lesage et al., 2016), whereas VPS13D is essential and thus at least one allele retains some VPS13D functions in affected individuals (Seong et al., 2018; Gauthier et al., 2018). As VPS13 proteins are ubiquitously expressed, it is unclear why mutations in each isoform result in distinct diseases. Human VPS13 proteins may have diverged to function at different subsets of MCSs. Indeed, VPS13A and VPS13C localize to contact sites that link the endoplasmic reticulum (ER) to mitochondria and endosomes respectively (Kumar et al., 2018), while VPS13B localizes to the Golgi and to endosomal vesicles (Seifert et al., 2011; Koike and Jahn, 2019). VPS13A and VPS13C are also found at ER-lipid droplet contact sites suggesting they may have both distinct and overlapping roles (Kumar et al., 2018; Yeshaw et al., 2019). Delineating the sites at which the different human paralogs function will provide a better understanding of how defects in lipid transport at specific MCSs contribute to neurological disorders. Because VPS13 is conserved in eukaryotes (Velayos-Baeza et al., 2004), much of our understanding of these proteins arises from studies performed in the budding yeast, Saccharomyces cerevisiae. Yeast has a single Vps13 protein that localizes to multiple organelle membranes and MCSs (Park and Neiman, 2012; Lang et al., 2015; John Peter et al., 2017; Park et al., 2016; Bean et al., 2018). Structural studies show that the highly conserved Vps13 N-terminus forms a hydrophobic groove and is sufficient for lipid transport between membranes in vitro (Kumar et al., 2018). This lipid transport function could explain why Vps13 can compensate for defects in the ER-mitochondrial encounter structure (ERMES), a lipid-transferring tether that forms ER-mitochondrial contact sites (Park et al., 2016; Lang et al., 57 2015). Other phenotypes of vps13Δ mutants, such as aberrant actin morphology and Golgi-endosome trafficking defects (Rzepnikowska et al., 2017; De et al., 2017), are not clearly linked to lipid transport although this may be because the responsible MCSs have not yet been identified. Understanding how VPS13 proteins are targeted to membranes could be key to understanding their role at different MCSs. We and others have recently demonstrated that specific adaptor proteins competitively target yeast Vps13 to different organelles (Bean et al., 2018; John Peter et al., 2017; Park et al., 2013) at different developmental stages or in response to environmental cues. The PX-domain protein Ypt35 recruits Vps13 to endosomes during exponential growth, while under starvation conditions, Ypt35 recruits Vps13 to the vacuole membrane at sites of contact with the nuclear ER (Bean et al., 2018). The mitochondrial adaptor, Mcp1, localizes Vps13 to the outer mitochondrial membrane (John Peter et al., 2017; Bean et al., 2018) whereas during meiosis, the sporulation-specific protein Spo71 recruits Vps13 to the prospore membrane (Park et al., 2013). These adaptors all possess a consensus PxP motif that interacts with Vps13 through a conserved domain found only in VPS13 proteins, which we have termed the Vps13 Adaptor Binding (VAB) domain (Bean et al., 2018). While the function of the VAB domain in human VPS13 proteins has not been determined, a VAB domain-containing fragment of VPS13C is sufficient for targeting to endolysosomal membranes suggesting that this domain may be important for the localization of at least some human VPS13 proteins (Kumar et al., 2018). Importantly, disease-causing missense mutations in VPS13B and VPS13D alter conserved asparagine residues in the VAB domain (Kolehmainen et al., 2004; Gauthier et al., 2018), suggesting it may have a conserved, functionally important role in yeast and humans. 58 The VAB domain consists of six repeats of approximately 100 residues with a highly conserved asparagine within the N-terminus of each repeat. It is not known if different adaptors bind different repeats and compete through steric hinderance, or if instead the VAB domain contains a single binding pocket that binds all motifs. In this study, we systematically mutate each of the yeast VAB domain repeats and demonstrate that the binding pocket for all known adaptors lies within repeats 5 to 6. We further demonstrate that a pathogenic missense mutation within the 6th VAB domain repeat of VPS13D blocks Vps13 adaptor binding and recruitment to membranes when modeled in yeast, suggesting the binding of VPS13D to adaptor proteins may be relevant for disease. 3.3 Results 3.3.1 Specific VAB domain mutations disrupt binding to the endosomal adaptor To determine the contribution of each repeat to adaptor binding, we generated a series of Vps13 point mutants with alanine substitutions in the highly conserved asparagine of each repeat (Bean et al., 2018) (Fig 3.1A). Herein these mutants are referred to as Vps13N1, Vps13N2, and so on, indicating the repeat with the substitution. Each mutant was tagged with GFP at an internal site that does not disrupt known Vps13 functions (Vps13^GFP) (Lang et al., 2015) and was expressed from a plasmid under its native promoter in a vps13∆ strain that co-expressed high levels of the HA-tagged endosomal adaptor, Ypt35. Mutation of the invariant asparagines did not alter protein levels substantially (Fig 3.1B), and mutations in repeats 2-5 did not significantly affect the co-immunoprecipitation (co-IP) of Ypt35 with Vps13 (Fig 3.1C,D). However, the Vps13N1 and Vps13N6 mutants demonstrated 59 Figure 3.1: Mutations in the Vps13 VAB domain disrupt its interaction with the endosomal adaptor Ypt35. (A) Schematic of Vps13 VAB domain indicating the position of the mutated conserved asparagines in repeats 1–6 (R1–R6). (B) Recovery of WT and mutant Vps13^GFP from anti-GFP immunoprecipitations analyzed by gel densitometry. One-way ANOVA with Dunnett-corrected post-hoc test; n=11; ***, P <0.001. (C) Co-IP of WT or mutant Vps13^GFP with Ypt35-3HA overexpressed from the ADH1 promoter. Proteins 60 were expressed on plasmids in vps13Δ strains. WCL, whole-cell lysate. (D) Densitometry analysis of Ypt35-3HA co-IP bands shown in C; one-way ANOVA with Dunnett-corrected post hoc test; n = 4; ***P < 0.001; **P < 0.01. (E) Localization of WT or mutant Vps13^GFP to bright puncta colocalizing with ADH1pr-driven Ypt35-RFP. Proteins were expressed on plasmids in ypt35Δ strains. (F) Automated quantitation of Vps13^GFP puncta from E. One-way ANOVA with Dunnett-corrected post hoc test; n = 3; cells/strain/replicate ≥1370; ****P < 0.0001; *P < 0.05. (G) Localization of WT or mutant Vps13^GFP to NVJ1-RFP-labeled NVJs in acetate-based media. Proteins were expressed on plasmids in vps13Δ strains. (H) Automated quantitation of the number of Vps13^GFP-labeled NVJs, from G. The diagram shows the location of the NVJ. One-way ANOVA with Dunnett-corrected post hoc test; n = 3, cells/strain/replicate ≥495; ****P < 0.0001; *P < 0.05. Error bars indicate SEM. Bars, 2 μm. strongly reduced pull-down of Ypt35, with respective recoveries of 24% (p<0.01) and 19% (p<0.001) relative to WT. We used fluorescence microscopy to determine the impact of each mutation on the Ypt35-mediated localization of Vps13^GFP to endosomes during exponential growth. In ypt35∆ strains expressing high levels of RFP-tagged Ypt35, wild type Vps13^GFP and most mutant forms localized to bright endosomal puncta that colocalized with Ypt35 (Fig 3.1E). Consistent with the decreased binding observed by co-IP, automated quantitation showed a near complete loss of localization of Vps13N1 and Vps13N6 mutants to puncta (Fig 3.1F; p<0.0001). A slight reduction in the punctate localization of Vps13N4 was also seen (70% relative to WT; p<0.05), although no significant loss of binding was detected by co-IP (Fig 3.1D). To examine if the Ypt35-mediated recruitment of Vps13 to nucleus-vacuole junctions (NVJs) was similarly affected, we analyzed the localization of the Vps13^GFP mutants in vps13∆ strains expressing the NVJ marker Nvj1-RFP. Upon glucose starvation, the Vps13N1 and Vps13N6 mutants remained cytosolic and did not colocalize with Nvj1-RFP (Fig 3.1G,H; p<0.0001). A minor defect was observed for Vps13N2, Vps13N4 and Vps13N5, which localized to 61 ~33% fewer NVJs compared to wildtype (Fig 3.1H; p<0.05). Taken together, these results suggest that the conserved asparagines of the 1st and 6th VAB domain repeats are necessary for the Vps13-Ypt35 interaction. 3.3.2 Similar VAB domain mutations disrupt the Vps13-Mcp1 interaction To determine if different VAB repeats are required for the interaction of Vps13 with the mitochondrial adaptor Mcp1, the Vps13^GFP mutants were immunoprecipitated from vps13∆ strains expressing high levels of HA-tagged Mcp1 (Fig 3.2A). As with Ypt35, mutation of the first and sixth repeats dramatically reduced the interaction with Mcp1 (24% and 6% recovery relative to WT respectively; p<0.0001). Modest reductions in Mcp1 recovery were seen for Vps13N3 (67%, p<0.05) and Vps13N4 (42%, p<0.0001) (Fig 3.2B). The recruitment of the Vps13^GFP mutants to mitochondria was examined in the strains overexpressing Mcp1, which enhances mitochondrial localization of Vps13 (John Peter et al., 2017; Bean et al., 2018) (Fig 3.2C). Automated analysis of the colocalization between Vps13^GFP and DAPI-stained mitochondrial DNA showed that Vps13N1 and Vps13N6 were defective in localizing to mitochondria (7% and 4% relative to WT; p<0.0001) (Fig 3.2D). In contrast, the mitochondrial localization of Vps13N4 was only slightly reduced (61% of WT) and that of Vps13N3 was not significantly altered, suggesting the modest reduction in Mcp1 binding is not sufficient to block Vps13 recruitment. Collectively, these results show that mutations in VAB domain repeats 1 and 6 strongly reduce binding of two different adaptors, Mcp1 and Ypt35. 62 Figure 3.2: VAB domain mutations disrupt Vps13 Mcp1-mediated mitochondrial localization. (A) Co-IP of ADH1pr-driven Mcp1-3HA with WT or mutant Vps13^GFP. WCL, whole-cell lysate. (B) Densitometry analysis of Mcp1-3HA co-IP bands shown in A; one-way ANOVA with Dunnett-corrected post hoc test; n = 4; ****P < 0.0001; *P < 0.05. (C) Colocalization of WT or mutant Vps13^GFP and DAPI-stained mtDNA in cells overexpressing Mcp1-3HA from an ADH1 promoter. (D) Automated quantitation of colocalized Vps13^GFP and mtDNA area/cell from C. One-way ANOVA with Dunnett-corrected post hoc test; n = 3, cells/strain/replicate ≥1012; ****P < 0.0001; **P < 0.01. Error bars indicate SEM. Bar, 2 μm 3.3.3 All three adaptors have similar requirements for VAB domain binding A similar strategy was used to determine if the interaction of Vps13 with the prospore membrane adaptor, Spo71, required the same VAB domain repeats as Ypt35 and Mcp1. When 63 Vps13^GFP mutants were immunoprecipitated in vps13∆ strains expressing high levels of the HA-tagged Spo71 (Fig 3.3A), only Vps13N1 and Vps13N6 had reduced pull-down of Spo71 (41% and 14% relative to WT; p<0.05 and p<0.01 respectively) (Fig 3.3B). During sporulation, Vps13 is recruited by Spo71 to the prospore membrane which elongates and closes around haploid daughter nuclei (Park et al., 2013). vps13∆ diploids have defects in prospore membrane expansion and closure preventing the formation of viable haploid gametes (Fig 3.3C) (Park and Neiman, 2012). To examine the phenotypic consequences of the reduced Vps13-Spo71 interaction, we determined the sporulation efficiency of vps13∆ diploids expressing each VAB domain mutant. All mutants gave rise to a similar number of viable haploids relative to wildtype, except for Vps13N6 which was nearly unable to form viable spores (p<0.0001) (Fig 3.3D). We were surprised that the Vps13N1 mutant, which showed a 60% reduction in Spo71 binding in our pull-down assay, did not have a sporulation defect. This suggests the level of Spo71 binding provided by Vps13N1 meets the threshold needed for wild type function. Taken together, our systematic mutational analysis of the Vps13 VAB domain shows that the binding of all adaptors is strongly affected by mutations in repeat 1 and 6, with repeat 6 being the most critical for adaptor interaction. 3.3.4 VAB domain mutations result in CPY secretion Vps13 was initially identified in a screen for mutants that missort the vacuolar hydrolase carboxypeptidase Y (CPY) (Bankaitis et al., 1986). In vps13∆ cells, defects in recycling the CPY receptor between the Golgi and endosomes leads to the entry of CPY into the secretory pathway and its subsequent release from the cell. Because the known adaptors are not required for this 64 Figure 3.3: VAB domain mutations disrupt Spo71-mediated sporulation and CPY sorting. (A) Co-IP of WT or mutant Vps13^GFP with Spo71-3HA overexpressed from a TEF1 promoter. WCL, whole-cell lysate. (B) Densitometry analysis of Spo71-3HA co-IP bands shown in A; one-way ANOVA with Dunnett-corrected post hoc test; n = 3; **P < 0.01; *P < 0.05. (C) Model of yeast sporulation depicting prospore membrane formation, elongation and closure. Red bars indicate steps in sporulation that are defective in vps13Δ strains. 65 (D) Mutation in the sixth VAB domain repeat prevents viable spore formation. Sporulated vps13Δ diploids carrying WT or mutant VPS13^GFP on plasmids were plated on selective media and colonies counted after 3 days of growth at 30°C. One-way ANOVA with Dunnett-corrected post hoc test; n = 3; ****P < 0.0001. (E) Mutations in the VAB domain result in CPY secretion. vps13Δ strains carrying WT or mutant VPS13^GFP on plasmids were spotted at the indicated dilutions, covered in nitrocellulose, grown overnight at 30°C and blotted with α-CPY antibody to detect secretion. Representative image shown; n = 3. (F) Overexpression of adaptor PxP-motifs results in CPY secretion that is rescued by mutating the conserved prolines within the motifs. The following fragments were expressed as RFP fusions from the strong TEF1 promoter: Ypt35(1–48), Mcp1(1–58) and Spo71(359–411) with PxP > AxA fragments containing Ypt35(P10,12A), Mcp1(P9,11A) and Spo71(P390,392A) substitutions. Error bars indicate SEM. Representative image shown; n = 3. process, we tested the effect of the VAB domain mutations on CPY sorting. Vps13^GFP mutants were spotted on plates and covered with a nitrocellulose filter to capture secreted CPY, which was detected by western blotting (Fig 3.3E). CPY was secreted by both Vps13N1 and Vps13N6 mutants, suggesting that the first and sixth repeats are necessary for the CPY sorting function of Vps13. This could indicate that a novel Vps13 adaptor is required for CPY sorting. We have previously shown that soluble chimeric proteins carrying a PxP motif compete with adaptors to block Vps13 recruitment to membranes (Bean et al., 2018). Therefore, if a new adaptor is required for Vps13’s CPY sorting function, overexpression of soluble PxP motif proteins should cause cells expressing wild type Vps13 to secrete CPY. Indeed, when the Ypt35 and Spo71 PxP motifs, but not the weaker Mcp1 motif, were fused to a cytosolic RFP marker and expressed from the strong TEF1 promoter in Vps13^GFP strains, a clear CPY sorting defect was observed that required the conserved prolines (Fig 3.3F). This suggests that Vps13 may be recruited by an as-yet-undiscovered PxP motif-containing adaptor to membranes where it functions in the CPY sorting pathway. 66 3.3.5 VAB repeats 5-6 are sufficient for adaptor binding Our results suggest that the integrity of the first and sixth repeats of the VAB domain are critical for binding all adaptor proteins. To identify the minimal subdomain required for adaptor binding and to rule out the possibility that these repeats form a single binding pocket in the folded protein, we expressed Envy-tagged forms of the first or last 3 VAB domain repeats from the ADH1 promoter in strains overexpressing HA-tagged Ypt35 (Fig 3.4A). Both 3 repeat-containing fragments were stably expressed but only the latter VAB repeats 4-6 bound Ypt35. This fragment pulled down all three adaptors in a PxP motif-dependent manner (Fig 3.4B-D). Further truncations of this minimal region determined that repeats 5-6, but not repeats 5 or 6 alone, could pull down Ypt35 (Fig 3.4E). Indeed, repeats 5-6 were sufficient for the PxP motif-dependent interaction of all 3 adaptors (Fig 3.4F-H). To determine if repeats 5-6 were sufficient for recruitment to membranes, we examined the localization of this fragment in ypt35Δ cells expressing high levels of RFP-tagged Ypt35 (Fig 3.4I). Indeed, GFP-tagged VAB repeats 5-6 colocalized with Ypt35-RFP labelled puncta, although fewer puncta were formed compared to full-length Vps13^GFP. Collectively, these results indicate that the adaptor binding pocket of the Vps13 VAB domain is located in repeats 5-6. This is consistent with our mutational analysis which showed that mutation of the invariant asparagine in repeat 6 severely reduced the binding of all three adaptors. In contrast, the conserved asparagine of repeat 1 appears to be important for binding only in the context of the full-length protein. Other repeats may also play a minor role in stabilizing the adaptor interaction or maintaining domain conformation. For example, mutations in repeats 3 and 4 reduced binding to the weakest affinity adaptor, Mcp1 (Fig 3.2A-B) (Bean et al., 2018), without significantly altering binding to Ypt35 or Spo71 (Fig 3.1C-D, Fig 3.3A-B). This 67 Figure 3.4: VAB domain repeats 5–6 are sufficient for binding all three adaptors. (A) VAB domain repeats 4–6 are sufficient for interaction with Ypt35. Co-IP of ADH1pr-driven Ypt35-3HA with ENVY-tagged Vps13 VAB domain truncations. 1–6* indicates an N-terminally extended VAB domain sequence. (B-D) Co-immunoprecipitations of overexpressed adaptors with WT or mutant PxP motifs, as indicated, and ADH1pr-driven ENVY-tagged Vps13 VAB repeats 4-6. Proteins were expressed on plasmids in vps13Δ strains. IP, immunoprecipitation; WCL, whole-cell lysate. Representative images shown; n = 3. (E) VAB domain repeats 5–6 are sufficient for interaction with Ypt35. Co-IP of Ypt35-3HA with ENVY-tagged Vps13 VAB domain repeats 4–6 truncations. (F–H) Adaptor interaction with VAB domain repeats 5–6 is PxP motif dependent. Co-IP of WT or PxP motif mutant overexpressed Ypt35-3HA, Mcp1-3HA and Spo71-3HA with ENVY-tagged Vps13 VAB repeats 5–6. WCL, whole-cell lysate. (I) GFP-tagged Vps13 VAB repeats 5–6 are sufficient for colocalization with ADH1pr-driven Ypt35-RFP. Representative images selected from n = 3. Bar, 2 μm. 68 suggests that weak affinity adaptors may provide a useful tool to identify subtle perturbations of the Vps13-adaptor interaction. 3.3.6 Human disease-causing mutations disrupt adaptor binding in yeast Vps13 Though the VAB domains of human VPS13 proteins lack known binding partners, disease-causing mutations in the VAB domains of VPS13B and VPS13D have been identified (Fig 3.5A). In VPS13D, a serine substitution in the conserved asparagine of VAB domain repeat 6, N3521S, is causative for spastic ataxia (Gauthier et al., 2018). In VPS13B, a serine substitution of the conserved asparagine of VAB domain repeat 4, N2993S, was identified as a homozygous mutation in a Belgian sibling pair presenting with Cohen syndrome (Kolehmainen et al., 2004). We modeled the VPS13D spastic ataxia mutation by mutating the corresponding N2428 residue in yeast Vps13^GFP to produce Vps13N6S and tested the effect on adaptor binding in a pull-down assay. The Vps13N6S mutant showed background level of binding to all three adaptors (n=3 per adaptor; Ypt35 = p<0.01; Mcp1 = p<0.0001; Spo71 = p<0.001), similar to the Vps13N6 mutant where the same residue was substituted with alanine (Fig 3.5B-E). This suggests the disease-causing mutation in VPS13D disrupts the function of the VAB domain in the context of yeast Vps13. We next modeled the pathogenic VPS13B N2993S mutation in yeast by mutating the corresponding N2216 residue in Vps13^GFP to create Vps14N4S. Vps13N4S had small reductions in the pull-down of Ypt35 and Mcp1, however only the reduction in Mcp1 pull-down was statistically significant (77% relative to wildtype; p<0.01) (Fig 3.5B-E), consistent with the effect of the Vps13N4A substitution (see Fig 3.2B). VPS13B has the least conserved VAB domain 69 Figure 3.5: Causative Cohen syndrome and spastic ataxia mutations result in perturbed adaptor binding in yeast. (A) Schematic of the VAB domains of VPS13B and VPS13D indicating the six repeats and position of conserved asparagines. VPS13B has a divergent VAB domain with serines in place of asparagines in repeats 2 and 3 indicated by red text. The causative Cohen syndrome mutation in VPS13B and spastic ataxia mutation in VPS13D are indicated in red boxes. (B–D) Co-IP of overexpressed Ypt35-3HA, Mcp1-3HA and Spo71-3HA with WT or mutant Vps13^GFP modeling the human mutations. WCL, whole-cell lysate. (E) Densitometry of immunoprecipitated HA-tagged adaptors from B–D. One-way ANOVA with Dunnett-corrected post hoc test; n = 3; ****P < 0.0001; ***P < 0.001; **P < 0.01; *P < 0.05. Error bars indicate SEM. of the human paralogs, with serines replacing the conserved asparagines in the 2nd and 3rd repeats. When these changes were modeled in yeast, creating Vps13N2,3S, we saw no effect on protein stability or adaptor pull-down (Fig 3.5B-E) suggesting these substitutions do not grossly disrupt the structural integrity of the VAB domain. However, more dramatic defects were seen when a N4S mutation was made in the context of this divergent VAB domain. The Vps13N2,3,4S 70 mutant showed a strongly decreased protein level, which likely accounts for the reduced pull-down of each adaptor (Fig 3.5B-D). While Ypt35 and Mcp1 were recovered at background levels (p<0.001 and p<0.0001 respectively), recovery of Spo71 was 42% relative to wildtype (p<0.05), suggesting the residual Vps13N2,3,4S was still capable of binding the highest affinity adaptor, Spo71 (Bean et al., 2018). These Vps13^GFP mutants were also examined for their ability to localize to organelles and mediate CPY sorting. Vps13N2,3,4S and Vps13N6S were not recruited to bright puncta by Ypt35-RFP (Fig 3.6A-B) (p<0.0001), or to mitochondria in cells expressing high levels of Mcp1 (Fig 3.6C-D) (p<0.01). Both Vps13N2,3,4S and Vps13N6S missorted CPY (Fig 3.6E), whereas no defect was seen for the Vps13N2,3S or Vps13N4S. Together these results suggest that VPS13B and VPS13D disease-causing mutations within the VAB domain result in reduced protein function when modeled in yeast, but for different reasons. Whereas the VPS13B mutation disrupts protein stability, the pathogenic VPS13D mutation does not significantly change protein levels, but instead may block its ability to interact with binding partners. 3.3.7 The VAB domain is comprised of six ß-sandwich folds After publication of the manuscript described in this chapter, the release of AlphaFold, an ab initio protein modeling software, allowed us to predict the structure of the VAB domain with high confidence (Jumper et al., 2021). Models of S. cerevisiae Vps13(1801-2650) generated with ColabFold (Mirdita et al., 2022) demonstrate that the VAB domain forms an arc-like structure comprised of six ß-sandwich folds with each repeat forming a single ß-sandwich (Fig 3.7A-C). The conserved asparagines of each repeat are at the end of a ß-sheet and face the interior of the 71 Figure 3.6: Causative Cohen syndrome and spastic ataxia mutations result in loss of yeast Vps13 localization and CPY secretion. (A) Disease-causing mutations in the Vps13 VAB domain block recruitment to ADH1pr-driven Ypt35-RFP. Proteins were expressed on plasmids in ypt35Δ cells. (B) Automated quantitation of Vps13^GFP puncta from A. One-way ANOVA with Dunnett-corrected post hoc test; n = 3, cells/strain/replicate ≥1164; ****P < 0.0001; **P < 0.01. (C) Disease-causing mutations in the Vps13 VAB domain block recruitment to mitochondria. Colocalization of Vps13^GFP mutants and DAPI-stained mtDNA in cells overexpressing Mcp1-3HA from an ADH1 promoter. (D) Automated quantitation of colocalized Vps13^GFP and mtDNA area/cell from C. One-way ANOVA with Dunnett-corrected post hoc test; n = 3, cells/strain/replicate ≥1200; ***P < 0.001; **P < 0.01. (E) Human disease-causing mutations in the VAB domain result in CPY secretion. vps13Δ cells carrying WT or mutant VPS13^GFP were blotted with α-CPY antibody to detect secretion as described in Figure 3.3E. Error bars indicate SEM. Bar, 2 μm. 72 Figure 3.7: AlphaFold modeling of the S. cerevisiae Vps13 VAB domain. (A) Top ranking model of Vps13(1801-2650) generated with ColabFold (Jumper et al., 2021; Mirdita et al., 2022) showing a structure comprised of repeating ß-sandwich folds. Residues are coloured according to their per-residue pLDDT values. (B) The per-residue pLDDT values for each of the 5 outputted ColabFold models. (C) Model of the VAB domain in A with individual VAB domain repeats coloured from red to purple. (D) The side chains of the conserved asparagines of each VAB domain repeat face the interior of a ß-sandwich. Vps13(2418-2548) encompassing VAB repeat 6 is shown with N2428 indicated. (E) ColabFold-generated predicted aligned error (PAE) plots for the Vps13(1801-2650) for rank 1, (F) rank 2, (G) rank 3, (H) rank 4, and (I) rank 5 models. The region encompassing the VAB domain is indicated in cyan. 73 ß-sandwich (Fig 3.7D), making it unlikely that these residues directly interact with adaptors and instead suggest a role in maintaining the domain conformation. The modeled Vps13 fragment included excess sequences on either end of the VAB domain which show this domain is connected to the hydrophobic lipid transfer channel of Vps13 through long unstructured loops that are modeled with low confidence (predicted local distance difference test (pLDDT) values ≤ 62) (Fig 3.7A-C). In our models, the lipid transfer segment appears in an orientation where the VAB domain could disrupt lipid flow through the remainder of this channel, however large predicted aligned error (PAE) values (>5 Å) between residues within the VAB domain and those within the channel suggest these regions are poorly modelled in relation to one another and could be oriented differently in the full-length protein (Fig 3.7E-I). Altogether, our modeling of this domain suggests that the Vps13 adaptor-binding pocket lies within the ß-sandwiches of repeats 5 and 6 and that mutation of the conserved asparagine of VAB repeat 6 likely disrupt the conformation of this pocket. 3.4 Discussion The VAB domain, which is found only in VPS13 proteins, is conserved in yeast and human isoforms suggesting it has an important functional role. Our studies of yeast Vps13 indicate that mutation of an invariant asparagine residue in the sixth repeat of this domain disrupts a critical protein interaction site. Because mutation of the corresponding residue in human VPS13D causes disease, both human and yeast VPS13 proteins may use this site to bind key regulators. This demonstrates that yeast Vps13 provides a useful model for studying the impact of human disease mutations. 74 3.4.1 The VAB domain is predicted to form an arc-like structure We found all adaptors respond similarly to alteration of the VAB domain repeats: mutation of the 6th repeat abolished the binding of all three adaptors tested, while mutation of the 1st repeat caused a partial defect. Moreover, we found that repeats 5 and 6 alone bound strongly to all three adaptors, suggesting adaptors recognize a single site within these repeats, consistent with our previous finding that the adaptors compete to recruit Vps13 to different membranes (Bean et al., 2018). Structural modeling predicts the VAB domain forms an arc-like structure where each repeat forms a β-sandwich fold. This domain appears to extend from the hydrophobic lipid transfer channel and could form the large arc observed on the otherwise rod-like reconstruction of yeast Vps13 obtained by negative-stain EM (De et al., 2017). The adaptor-binding site lies within the last two β-sandwiches of the VAB domain, and because mutation of the conserved asparagine in the 6th repeat nearly abolished adaptor binding, whereas mutation of the analogous residue in the 5th repeat had no effect, it is tempting to conclude that the “N6” residue lies near the binding pocket. As the conserved asparagine of each repeat faces the interior of a β-sandwich, it is unlikely that the repeat 6 asparagine interacts directly with the PxP peptide, but instead likely helps maintain the proper conformation of the binding site by stabilizing the interfaces between β-sheets or by maintaining the position of interacting loops. Mutation of the conserved asparagine in the 1st VAB domain repeat also had a pronounced effect on adaptor binding. However, repeat 1 was dispensable for binding when truncated fragments of the VAB domain were expressed. Repeat 1 could contribute indirectly to adaptor binding by interacting with an additional cofactor at membranes. Mutations in repeat 1 that block the interaction with putative regulators could thus prevent adaptor binding to repeats 75 5-6. It is also possible that this mutation more generally disrupts the orientation of the VAB domain at membranes due to misfolding proximal to the site of attachment to the hydrophobic lipid transfer channel, which could prevent interactions with membrane-associated adaptors. The C-terminal location of the Vps13 VAB domain suggests the adaptor-binding site may be kept close to one opposing membrane at contact sites, and thus adaptor interactions could specify the donor or acceptor membranes for lipid transfer. This model for MCS recruitment is similar to that of the related ATG2 proteins (Fig 3.8). Although ATG2 proteins lacks a VAB domain, C-terminal interactions with the phosphatidylinositol 3-phosphate (PI(3)P)-binding WD40 protein WIPI4/WDR45 (Atg18 in yeast) are necessary to enhance human ATG2A/B recruitment to isolation membranes for their function in autophagosome elongation and closure (Chowdhury et al., 2018; Watanabe et al., 2012; Baskaran et al., 2012). Additional conserved C-terminal regions such as amphipathic helices in Atg2 (Tamura et al., 2017; Kotani et al., 2018a) and Vps13 (Kumar et al., 2018), or the predicted PH domain in Vps13 (Fidler et al., 2016), may help stabilize recruitment through coincidence detection. Further work will be needed to determine the role of these additional domains in targeting Vps13-like proteins to contact sites, and structural studies will be needed to verify the VAB domain structure and identify the key residues involved in the Vps13-adaptor interaction. 3.4.2 New adaptors in yeast and humans Our work suggests the full complement of adaptor proteins that bind the yeast Vps13 VAB domain has not yet been uncovered and that an undiscovered PxP-containing adaptor likely regulates Vps13’s CPY sorting function. Such an adaptor may recruit Vps13 to Golgi membranes to function in Golgi/endosome transport and homotypic fusion (De et al., 2017). 76 Figure 3.8: Model of related membrane recruitment strategies of Vps13 and Atg2 proteins. Vps13 and Atg2 are recruited to membranes through adaptor-mediated interactions. In Vps13, adaptor proteins bind within VAB domain repeats 5-6 through related PxP motifs to recruit Vps13 to different organelle membranes. In Atg2, recruitment to isolation membranes is mediated by interactions with the 7-bladed WD-40 protein, WIPI4/WDR45 in humans or Atg18 in yeast, which bind PI(3)P (Watanabe et al., 2012; Baskaran et al., 2012). Other conserved C-terminal domains, such as the helical ATG2_C domains and a predicted PH domain, may help position these proteins for efficient lipid transfer. Arrows indicate the putative direction of lipid transport along a hydrophobic groove shown by dotted lines. The N- and C-termini of both proteins are indicated. Vps13 has also been localized to other organelles, including peroxisomes (John Peter et al., 2017), which could require yet other adaptors. These unidentified adaptors may be discovered by bioinformatics approaches that identify predicted PxP motifs (Bean et al., 2018). Novel adaptor proteins may also bind the VAB domains of human VPS13 proteins. The dramatic effect of the disease-causing VPS13D mutation when modeled in yeast strongly suggests that VPS13D has a conserved adaptor binding site. However, the PxP motif is not conserved beyond the order of Saccharomycetales suggesting the yeast PxP motif co-evolved 77 with the VAB domain. It is interesting to speculate that the human VPS13 proteins have diverged to recognize distinct motifs that target them to discrete MCSs. 3.4.3 The role of the VAB domain in human VPS13 proteins VPS13D is an essential gene and thus disease-causing mutations are typically compound heterozygous with one null and one missense VPS13D allele (Seong et al., 2018; Gauthier et al., 2018). Because the VPS13DN6S mutation is paired with a frame-shift mutation that removes nearly half of the protein sequence (Gauthier et al., 2018), it is likely that the VAB domain mutation maintains essential VPS13D functions while blocking a specific adaptor interaction that results in ataxia. Two other disease-causing missense mutations in VPS13D fall within the VAB domain in repeat 1 (L2900S) and repeat 3 (R3253N) (Gauthier et al., 2018). The effect of these mutations has not been studied, but the clustering of causative missense mutations in the VPS13D VAB domain is consistent with an important role in disease for this domain. Although the localization of VPS13D is unknown, the abnormal mitochondrial morphology seen in affected patient fibroblasts and in Drosophila models suggests that VPS13D may function at mitochondria (Seong et al., 2018; Anding et al., 2018). If human VPS13 proteins have evolved different localization mechanisms to target distinct contact sites, the VAB domains of some paralogs may have lost their adaptor binding function. This could be true for VPS13B, which has a poorly conserved VAB domain and is recruited to Golgi membranes through interactions with Rab6 within its C-terminus (Seifert et al., 2015). The divergent VAB domain of VPS13B has serine residues in the place of conserved asparagines in the 2nd and 3rd VAB domain repeats. Modeling these substitutions in yeast suggests they have little or no effect on protein function, and naturally occurring asparagine-to-78 serine substitutions in VAB repeats 3 and 5 are also present in members of the division Magnoliaphyta and Ascomycota, respectively. The observation that this divergent VAB domain is destabilized by an asparagine-to-serine substitution introduced into repeat 4 could explain why the N2993S mutation in VPS13B causes Cohen syndrome, which is typically characterized by a loss of VPS13B protein (Balikova et al., 2009; Seifert et al., 2006). These results demonstrate the utility of the yeast system for modeling human disease mutations. Further studies in yeast are expected to generate new insights into membrane targeting mechanisms of the VPS13 family and will provide a useful system for testing the effects of candidate pathogenic mutations. Do other VPS13 VAB domains function in protein localization? A fragment of VPS13C containing its VAB domain is sufficient for localization to endolysosomal membranes (Kumar et al., 2018). A similar VPS13A fragment was found to be cytosolic, however a missense mutation in this domain results in chorea-acanthocytosis (Velayos-Baeza et al., 2008). This suggests that VPS13A may rely on the VAB domain for localization, although the relevant adaptors may only be present in certain cell types, developmental stages or conditions. Other features of the VPS13 proteins may work together for targeting to membrane contact sites. VPS13A and VPS13C both have additional localization determinants including a FFAT motif for ER targeting and an amphipathic helix for targeting to lipid droplets (Kumar et al., 2018). Additionally, all human VPS13 proteins have been found to bind Rab proteins in high and low-throughput studies (Koike and Jahn, 2019; Seifert et al., 2015; Gillingham et al., 2014). Other regions of Vps13 may interact with lipids to stabilize membrane recruitment through coincidence detection. Yeast Vps13 binds lipids through regions at its N- and C-termini, and a causative chorea-acanthocytosis mutation in the APT1 domain disrupts binding to PI(3)P when modeled in yeast (De et al., 2017; Rzepnikowska et al., 2017). Identifying the different targeting 79 mechanisms used by each human VPS13 protein will be important for determining the subset of MCSs where each paralog functions and for unravelling the role of specific MCSs in disease. 3.5 Materials and methods 3.5.1 Yeast strains and plasmids Strains and plasmids used in this study are described in Table 3.1 and Table 3.2 respectively. All strains from this study were made by PCR-based homologous recombination as described (Longtine et al., 1998; Janke et al., 2004). Gene deletions were confirmed by colony PCR. Plasmids were made by homologous recombination in yeast by co-transforming linearized plasmids with generated PCR products. The plasmids were recovered in Escherichia coli and sequenced. Table 3.1: Yeast strains used in Chapter 3 Strain ID Genotype Source BY4741 vps13Δ BY4741 MATa vps13Δ::kanMX4 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Giaever et al., 2002) MDY1258 BY4741 MATa ypt35Δ::kanMX4 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Bean et al., 2018) KK01 BY4741 MATa vps13Δ::kanMX4 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Dziurdzik et al., 2020) MDY293 Y7043 MATα vps13Δ::natMX4 his3Δ1 met15Δ0 ura3Δ0 can1Δ::STE2pr-LEU2 lyp1Δ cyh2Δ LYS2+ (Dziurdzik et al., 2020) ByK337 BY4741 MATa VPS13^499GFP his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Lang et al., 2015) KK30 BY4742 MATa vps13Δ::kanMX4 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Dziurdzik et al., 2020) KK17 BY4742 MATα kanMX4::ADH1pr-VPS13-GFP+::HIS3 leu2Δ0 met15Δ0 ura3Δ0 (Bean et al., 2018) BY4742 BY4742 MATα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 Dr. E.K. O’Shea (Harvard University) 80 Table 3.2: Plasmids used in Chapter 3 Plasmid ID Plasmid Description Source pRS416 pRS416 (URA3, CEN) ATCC #87521 pMD232 pRS415-ADH1pr-YPT35-3HA (LEU2, CEN) (Bean et al., 2018) pMD218 pRS416-VPS13^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD312 pRS416-VPS13(N1875A (N1))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD317 pRS416-VPS13(N1991A (N2))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD325 pRS416-VPS13(N2108A (N3))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD313 pRS416-VPS13(N2216A (N4))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD316 pRS416-VPS13(N2326A (N5))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD314 pRS416-VPS13(N2428A (N6))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD195 pRS415-ADH1pr-YPT35-RUBY2 (LEU2, CEN) (Bean et al., 2018) pBB40 pRS415-ADH1pr-MCP1-3HA (LEU2, CEN) (Bean et al., 2018) pMD166 pRS425-TEF1pr-SPO71-HA (LEU2, 2μ) (Bean et al., 2018) pBB73 pRS416-TEF1pr-mCherry (URA3, CEN) (Bean et al., 2018) pBB66 pRS416-TEF1pr-YPT35(1-48)-mCherry (URA3, CEN) (Bean et al., 2018) pBB74 pRS416-TEF1pr-YPT35(1-48; P10,12A)-mCherry (URA3, CEN) (Bean et al., 2018) pBB67 pRS416-TEF1pr-MCP1(1-58)-mCherry (URA3, CEN) (Bean et al., 2018) pBB75 pRS416-TEF1pr-MCP1(1-58; P9,11A)-mCherry (URA3, CEN) (Bean et al., 2018) pBB65 pRS416-TEF1pr-SPO71(359-411)-mCherry (URA3, CEN) (Bean et al., 2018) pBB76 pRS416-TEF1pr-SPO71(359-411; P390, 392A)-mCherry (URA3, CEN) (Bean et al., 2018) pMD290 pRS416-ADH1pr-VPS13(1732-2556)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD385 pRS416-ADH1pr-VPS13(1821-2556)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD386 pRS416-ADH1pr-VPS13(1821-2199)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD384 pRS416-ADH1pr-VPS13(2201-2556)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD387 pRS416-ADH1pr-VPS13(1821-1967)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD388 pRS416-ADH1pr-VPS13(2409-2556)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD291 pRS415-ADH1pr-YPT35(P10,12A)-3HA (LEU2, CEN) (Bean et al., 2018) pBB63 pRS415-ADH1pr-MCP1(P9,11A)-3HA (LEU2, CEN) (Dziurdzik et al., 2020) pBB59 pRS425-TEF1pr-SPO71(P390,392A)-3HA (LEU2, 2μ) (Bean et al., 2018) pSD033 pRS416-ADH1pr-VPS13(2201-2409)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) 81 Plasmid ID Plasmid Description Source pMD400 pRS416-ADH1pr-VPS13(2306-2556)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pSD032 pRS416-ADH1pr-VPS13(2201-2306)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pSD031 pRS416-ADH1pr-VPS13(2306-2409)-ENVY (URA3, CEN) (Dziurdzik et al., 2020) pMD322 pRS416-VPS13(N2216S (N4S))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD383 pRS416-VPS13(N1991,2108S (N2,3S))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD382 pRS416-VPS13(N1991,2108,2216S (N2,3,4S))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pMD351 pRS416-VPS13(N2428S (N6S))^GFP (URA3, CEN) (Dziurdzik et al., 2020) pNVJ1-RFP pRS415-NVJ1-mCherry (LEU2, CEN) Dr. M. Henne (University of Texas Southwestern Medical Center) 3.5.2 Co-immunoprecipitations For Vps13^GFP point mutants, yeast were grown to log-phase in minimal synthetic dextrose media and 60 OD600 of cells stored at -80 °C until further use. Cells were thawed and resuspended in 500 µL lysis buffer (50 mM HEPES, 0.1% Tween-20, 50 mM NaCl, 1 mM EDTA, 1 mM PMSF, and 1× yeast/fungal ProteaseArrest, pH 7.4) and lysed by vortexing with 0.5 mm diameter glass beads. 50 µL of lysate was set aside and resuspended in 4× Laemmli Sample Buffer with 8M urea (8% SDS, 40% glycerol, 240 mM Tris-Cl pH 6.8, 0.004 % bromophenol blue, 20% β-mercaptoethanol). The remainder of the lysates were incubated with polyclonal rabbit anti-GFP (EU2; Eusera) for 1 hour followed by Protein A-Sepharose beads (GE Healthcare) for another hour at 4° C. Washed beads were resuspended in 50 µL of Thorner buffer (8 M urea, 5% SDS, 40 mM Tris pH 6.8, 0.1 M EDTA, 1% β-mercaptoethanol, 0.4 mg/mL bromophenol blue) and proteins eluted by heating at 100° C for 5 minutes. Samples were run on SDS-PAGE gels, transferred to nitrocellulose membranes and blotted with either polyclonal rabbit anti-GFP (EU2; Eusera) and Tidyblot Western Blot Detection Reagent:HRP 82 (STAR209P; Bio-Rad) or monoclonal mouse anti-HA (MMS-101R; Covance) and horseradish peroxidase-conjugated polyclonal goat anti-mouse (115-035-146; Jackson ImmunoResearch Laboratories). For co-immunoprecipitations with Vps13 VAB domain truncations, 50 OD600 of yeast were lysed. The nitrocellulose membranes were blotted with either monoclonal mouse anti-GFP (11-814-460-001; Roche) or monoclonal mouse anti-HA (MMS-101R; Covance) followed by horseradish peroxidase-conjugated polyclonal goat anti-mouse (115-035-146; Jackson ImmunoResearch Laboratories). All blots were developed with West Pico (Pierce) or Amersham ECL Prime (GE Healthcare) chemiluminescent reagents and exposed to Amersham Hyperfilm (GE Healthcare). Films were scanned, and densitometry performed in Fiji (Schindelin et al., 2012). 3.5.3 Fluorescence microscopy and automated quantitation Yeast were grown to log-phase in minimal synthetic dextrose media, transferred to concanavalin A-treated glass bottom cell imaging plates (Eppendorf) and imaged on a DMi8 microscope (Leica Microsystems) with a high-contrast Plan Apochromat 63×/1.30 Glyc CORR CS objective (Leica Microsystems), an ORCA-Flash4.0 digital camera (Hamamatsu Photonics), and MetaMorph 7.8 software (MDS Analytical Technologies). Mitochondrial DNA was labelled with 1 µg/mL DAPI (Sigma-Aldrich) for 20 minutes at 30° C. Cells were washed prior to imaging. For NVJ localization, 2 OD600/mL of log-phase yeast were incubated at 30° C for 18 hours in 2 mL synthetic complete media with 2% acetate as the carbon source. Yeast were imaged as above except washed cells were maintained in nutrient-depleted synthetic acetate media during imaging. 83 All measurements were done on raw images; for presentation, images were processed with MetaMorph 7.8 software. Cropping and adjustment of brightness and contrast was performed in Photoshop CC 2017 (Adobe). Images were analyzed using custom MetaMorph 7.8 journals. Live cells were identified through several iterations of the Count Nuclei function. Puncta number and size were identified with the Granularity function. Puncta attributed to dead cells and extracellular noise were masked by using the LogicalAND function. The area of colocalization of Vps13^GFP and DAPI-labelled mitochondria, identified with the Granularity function, were determined by using the built-in Measure Colocalization application. The colocalizing Vps13^GFP and NVJ1-RFP structures were determined by masking NVJs identified by the TopHat function in both channels and counted using the Granularity function. 3.5.4 CPY secretion To examine the secretion of CPY, 1 OD600 of log-phase yeast were serially diluted and spotted on minimal synthetic dextrose media plates. The plates were covered with nitrocellulose and incubated overnight at 30° C. The nitrocellulose membranes were washed to remove cells and blotted with monoclonal mouse anti-carboxypeptidase Y (A-6428; Thermo Fisher Scientific). Blots were developed with West Femto (Pierce) chemiluminescent reagent and exposed to Amersham Hyperfilm (GE Healthcare). 3.5.5 Sporulation To examine sporulation efficiency, 0.25 OD600 of log-phase diploid yeast were inoculated in 5 mL sporulation media (0.3% Kac, 0.2% raffinose, 0.012% amino acid mix) and incubated at 84 room temperature for 4 days. The cells were washed, diluted and plated on selective media to select for spores. Plates were incubated at 30° C and colonies counted after 3 days. 3.5.6 Statistical analysis Statistical analyses were carried out by performing one-way ANOVA with a Dunnett-corrected post hoc test for comparisons to WT using Prism 6 (GraphPad Software). Data distribution was assumed to be normal, but this was not formally tested. 3.5.7 Protein structure prediction Ab-initio prediction of the VAB domain protein structure was performed by the AlphaFold2 Mmseq2-driven homology search software ColabFold 1.3.0 (Mirdita et al., 2022; https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb) with default settings. Figures were generated in UCSF Chimera version 1.16.0 (Pettersen et al., 2004; University of California San Francisco Parnassus Campus) and assembled in Adobe Illustrator 2022 (Adobe, Mountain View, California). 85 Chapter 4: A conserved interactor targets Fmp27 to ER-PM contact sites for a shared role in lipid homeostasis 4.1 Introduction Eukaryotic cells are compartmentalized into membrane-bound organelles that have distinct lipid and protein compositions required for their unique functions. The majority of cellular lipids are synthesized in the endoplasmic reticulum (ER) and then transported to other organelles through both vesicular and non-vesicular trafficking pathways (van Meer et al., 2008; Stefan et al., 2017). In non-vesicular lipid transport, specialized lipid transfer proteins (LTPs) move lipids between organelles at membrane contact sites (MCSs), regions where organelles are tethered together in close proximity (Prinz et al., 2020; Scorrano et al., 2019; Wong et al., 2019). LTPs are particularly important for lipid transport to organelles that are not connected to the vesicular transport pathway, such as mitochondria and peroxisomes. Additionally, unlike vesicular transport where heterogenous bulk lipids are simultaneously trafficked, LTPs transport specific lipid species allowing for maintenance and rapid alterations of organelle lipid composition in response to environmental cues or stresses (Prinz et al., 2020; Wong et al., 2019; Kim et al., 2022) To date, LTPs have been shown to mediate lipid transfer at MCSs either through shuttling or bridging mechanisms. The majority of the previously characterized LTPs shuttle lipids between donor and acceptor membranes using specialized domains that consist of a hydrophobic cavity that can bind individual lipid moieties and shield them from the aqueous cytosol during transfer (Wong et al., 2019; Reinisch and Prinz, 2021). One well characterized example of shuttling LTPs is the Lipid transfer proteins Anchored at Membrane contact sites 86 (LAM) family which transport sterols at contacts sites via Steroidogenic Acute Regulatory Transfer (StART) domains (Jentsch et al., 2018; Tong et al., 2018; Besprozvannaya et al., 2018). More recently, a novel class of bridging LTPs has been identified. Members of this class form elongated structures that can tether organelles at either end of a MCS and transport lipids directly between them (Wong et al., 2019; Reinisch and Prinz, 2021; Neuman et al., 2022b). Vps13 and Atg2 were the first members of this class to have a portion of their lipid transfer module structurally resolved. Both proteins have conserved N-terminal chorein domains that form an elongated groove with a hydrophobic interior that can bind numerous lipids (Kumar et al., 2018; Osawa et al., 2019). Cryo-EM structures of full-length or extended portions of these proteins suggest this hydrophobic groove structure, comprised of b-sheets with overarching a-helices, extends throughout most of the protein forming a continuous lipid transfer channel (Li et al., 2020; Adlakha et al., 2022; De et al., 2017; Chowdhury et al., 2018; Maeda et al., 2019). Due to this Repeating Beta Groove (RBG) domain, this family of bridging LTPs has tentatively been named the RBG superfamily (Levine, 2022; Neuman et al., 2022b). The elongated hydrophobic channels of RBG proteins could allow lipids to rapidly slide across a contact site from one membrane to another, and thus RBG proteins may function in membrane expansion events where bulk lipids must rapidly be supplied, such as during sporulation (Vps13) or autophagy (Atg2) (Park and Neiman, 2012; Velikkakath et al., 2012; Valverde et al., 2019). Additional RBG superfamily LTPs have recently been identified, however the full repertoire of related proteins in this family is not known. Newly identified members include mammalian BLTP3B/SHIP164, and its paralog BLTP3A/UHRF1BP1, which function in retrograde trafficking from endosomes to Golgi (Hanna et al., 2022), yeast Csf1 and its nematode homolog, LPD-3, which maintain phospholipid distribution at ER-plasma membrane 87 (PM) contacts and function in cold tolerance (John Peter et al., 2022; Toulmay et al., 2022; Wang et al., 2022), and fly Hobbit which is required for regulated secretion events (Neuman and Bashirullah, 2018). It is unclear whether these newly identified LTPs also function in membrane expansion events, or whether they function in broad organelle lipid homeostasis. Hobbit has two orthologs in budding yeast, Fmp27/Hob1 and Ypr117w/Hob2, which have similarly been shown to localize to ER-PM contacts like their fly counterpart (Neuman et al., 2022a; Toulmay et al., 2022), but to date no function has been determined. To operate as lipid transport bridges, Fmp27 and Hob2 must target membranes on either end of a contact site. Fmp27 has been shown to target the ER through its N-terminus, similar to Vps13 and Atg2 (Kotani et al., 2018a; Neuman et al., 2022a). The PM targeting mechanism for the Fmp27 C-terminus is unknown, but could rely on interactions with organelle-specific adaptors as is seen for other RBG members (Kotani et al., 2018a; Maeda et al., 2019; Gómez-Sánchez et al., 2018; Park et al., 2013; John Peter et al., 2017; Bean et al., 2018; Dziurdzik et al., 2020). Both the role and targeting mechanism for Fmp27 and Hob2 paralogs at ER-PM contact sites are unknown. Here we determine that Fmp27 is targeted to the PM through interactions with the previously uncharacterized Ybl086c, which we have named Hoi1 for Hobbit Interactor 1. Using ColabFold, an AlphaFold2-based protein structure prediction software, we identify a key interaction interface at an ⍺-helical extension conserved in both Fmp27 and Hob2. We demonstrate that loss of both paralogous proteins, or their PM interactor, result in enhanced sensitivity to sterol-binding drugs and a redistribution of accessible sterols from the cytosolic PM leaflet to intracellular puncta, and place these proteins in a shared lipid homeostasis pathway together with the Lam protein family. 88 4.2 Results 4.2.1 Fmp27 and Hob2 are RBG proteins with homology to Vps13 When the work described in this chapter was begun, no other members of the RBG superfamily besides Vps13 and Atg2 were known. To uncover putative novel yeast protein family members, we used HHpred, a remote protein homology detection server (Söding et al., 2005), to identify proteins with predicted structural homology to the Vps13 lipid transfer channel (Fig 4.1A). As query, we used the sequence of a central portion of Vps13 (residues 496-1390) that maps to the most highly resolved portion of the channel (<4.5 Å resolution; Li et al., 2020), and that we found modeled the characteristic RBG domains with the trRosetta server (Du et al., 2021). This HHpred analysis identified 5 homologous proteins: Vps13, Atg2, Fmp27/Hob1, Ypr117w/Hob2 and Csf1. Unexpectedly, the remote homologies identified in Atg2, Fmp27 and Hob2 mapped to annotated APT1 domains, a domain also present in Vps13, which has been proposed to function as a phospholipid-binding localization determinant (Rzepnikowska et al., 2017; Kolakowski et al., 2020). To determine if the Vps13 APT1 domain has structural homology to the lipid transport channel, a reciprocal HHpred analysis was performed with this domain as query (Fig 4.1B). The analysis indeed identified the same homologous yeast proteins and regions, with the exception of Csf1, suggesting the APT1 domain is a continuation of the lipid transport channel. To assess this further, the Vps13 protein sequence was subdivided into 250 residue non-overlapping sections and direct comparisons of all the fragments against one another were performed with HHpred (Fig 4.1C). In support that this analysis can identify regions of homology, the 3 fragments that contain the six-repeat Vps13 Adaptor Binding (VAB) domain (Dziurdzik et al., 2020; Adlakha et al., 2022) were identified as homologous with true positive probabilities ≥97%. Further, as 89 Figure 4.1: HHpred analysis identifies predicted structural homologs of Vps13. HHpred analysis of the Saccharomyces cerevisiae proteome identifies predicted structural homologs of Vps13 using (A) a portion of the elongated lipid transport channel (residues 496-1390) or (B) the annotated APT1 domain (residues 2575-2832) as queries. The homologous regions identified are shown with their true positive probabilities indicated; red ≥ 90%; orange ≥85% and yellow <85%. The annotated domains are: TM = predicted transmembrane domain; Chorein domain; Vps13 Adaptor Binding (VAB) domain, aberrant pollen transmission 1 (APT1) domain; Autophagy-related protein 2 C-terminal (ATG2_C) domain; and Pleckstrin homology (PH) domain. The PDB_mmCIF70_3_Mar structural/domain database and Euk_Saccharomyce_cerevisiae_S288C_11_ 90 Mar_2017 proteome were used. (C) Matrix presenting the maximum true positive probability for predicted structural homology from comparative HHpred analysis of all non-overlapping 250 amino acid regions of Vps13. The comparisons are coloured according to their true positive probabilities with higher probabilities shown in darker green. Comparisons of identical fragments are in gray. The analysis identifies structural homology within the elongated lipid channel and the APT1 domain; as well as separate homology around the six-repeat Vps13 Adaptor Binding (VAB) domain. Domain schematics of Vps13 are aligned to the right and bottom of the matrix as a guide. Domains are coloured according to the schematics in A and B. (D) Predicted structural models of the annotated APT1 domains of Fmp27 and Hob2 highlighting the similar ß-sheet elongated groove structure seen in Vps13 and Atg2 proteins. Models are coloured by per-residue pLDDT values. Structures were extracted from full-length structural predictions acquired from the AlphaFold Protein Structure Database (https://alphafold.ebi.ac.uk/; Jumper et al., 2021). (E) Structural predictions of paralogous Fmp27 and Hob2 proteins acquired from the AlphaFold Protein Structure Database. Models are coloured by per-residue pLDDT values. Both proteins form elongated channels comprised of repeating ß-sheets and a similar arrangement of ⍺-helical protrusions. Bar = 10nm. expected, the regions containing the elongated lipid transport channel but excluding the chorein domain (residues 251-1750) had high true positive probability scores when compared to each other. These regions also identified homology to the APT1 domain with true positive probabilities ≥89%, supporting the hypothesis that the APT1 domain is an extension of the N-terminal lipid transport channel (Adlakha et al., 2022). To validate the structural similarity of the identified Vps13 homologs, we focused on the paralogs Fmp27 and Hob2, high confidence hits in both HHpred searches, that are conserved in metazoans. The advent of the ab initio protein modeling software, AlphaFold (Jumper et al., 2021),which was released to the public after our HHpred analysis was completed, allowed the large full-length proteins to confidently be modeled in their entirety. Initial inspection of the AlphaFold-predicted structures determined that the annotated APT1 domains of Fmp27 and Hob2 fold into the characteristic repeating ß-sheet groove with overarching ⍺-helices (Fig 4.1D), resembling the solved portions of the Vps13 lipid transfer channel (Kumar et al., 2018; Li et al., 91 2020). Further inspection of the full-length models determined that both Fmp27 and Hob2 are structurally related proteins with elongated hydrophobic channels consisting of tandem RBG domains and a similar orientation of a-helices (Fig 4.1E). The predicted channels of both Fmp27 and Hob2 extend to ~30nm, a distance that would permit these paralogs to span a contact site and bridge membranes on either side. In parallel, others have used a similar approach to independently identify Fmp27, Hob2 and Csf1 as structurally homologous proteins to Vps13 and Atg2 (Castro et al., 2022; John Peter et al., 2022; Toulmay et al., 2022; Neuman et al., 2022b). Taken together, this strongly suggests that Fmp27 and Hob2 are related members of the RBG superfamily. 4.2.2 Fmp27 localizes to ER-PM contact sites Recent studies have shown that C-terminally tagged forms of Fmp27 and Hob2 localize to ER-PM contact sites (Neuman et al., 2022a; Toulmay et al., 2022). However, tags at either N- or C-termini of Vps13 interfere with its localization and function (Lang et al., 2015), and similar tag interference may perturb the localization of Fmp27 and Hob2 (Castro et al., 2022). To validate the endogenous localization of Fmp27, we designed a yeast strain with the fluorophore NeonGreen (NG) inserted in an unconserved loop after residue 737 (Fmp27^NG). We determined that Fmp27^NG, like Fmp27-ENVY, localizes to cortical sites that colocalize with an RFP-labelled ER marker, Sec63-mScarletI, and partially colocalizes with an ER-PM contact site protein, TCB3-mScarletI (Fig 4.2A). Due to the low expression of Hob2 and poor visualization, we decided to focus our further characterization on Fmp27. The partial colocalization of Fmp27 with Tcb3 suggests that they may preferentially reside at distinct ER-PM contact sites. Notably, Tcb3 has been reported to localize to PM 92 Figure 4.2: Fmp27 resides at ER-PM contact sites. (A) Confocal images of live yeast cells expressing Fmp27^NeonGreen (NG) and Sec63-mScarletI, an ER marker, or TCB3-mScarletI, an ER-PM contact site marker. All proteins are expressed from their endogenous genomic loci. Linearized line scans of the cell cortex and normalized fluorescence signal intensities are shown to the right with curved arrows in the merged image indicating the line scan start site. Yellow arrowheads indicate sites of colocalization. Bar = 2µm. (B) Fmp27-ENVY maintains colocalization with SEC63-mScarletI in rtn1Δ yop1Δ cells. Fluorescent images of wild-type and rtn1Δ yop1Δ yeast cells expressing Sec63-mScarletI and either Tcb3-EGFP, sfGFP-Scs2 or Fmp27-ENVY. All proteins are expressed from their endogenous genomic loci. Linearized line scans of the cell cortex and normalized fluorescence intensities are shown to the right with curved arrows in the merged image indicating the line scan start site. Bar = 2µm. 93 contacts with tubular ER with high membrane curvature (Hoffmann et al., 2019; Collado et al., 2019). This ER-PM contact site can be discriminated from PM contacts with flat ER sheets in rtn1Δ yop1Δ mutants that lack 2 of the 3 reticulon proteins that mediate ER tubulation (Hoffmann et al., 2019). In this mutant, ER-PM contact site proteins that localize preferentially to ER sheets, such as Scs2, maintain colocalization with the ER translocon Sec63, while those at residual curved ER lose colocalization (Hoffmann et al., 2019). Thus, we assessed the correspondence of Fmp27-ENVY signals with Sec63-mScarletI at the cell cortex, along with those of Tcb3-EGFP or sfGFP-Scs2 in wild type (WT) and rtn1Δ yop1Δ mutants. Fmp27-ENVY, like sfGFP-Scs2, was found to colocalize with Sec63-mScarletI in both WT and rtn1Δ yop1Δ mutant yeast, while in contrast Tcb3-EGFP lost colocalization in the mutant (Fig 4.2B). This supports a preference for different ER-PM contact sites between Fmp27 and Tcb3, and further suggests that Fmp27 resides at PM contacts with flat ER sheets. To function as a lipid transport channel at contact sites, both termini of Fmp27 must target opposing organelle membranes. A predicted N-terminal anchor is responsible for targeting Fmp27 to the ER. Further, FMP27 overexpression results in a redistribution of Fmp27 along the entire ER from discrete cortical ER-PM puncta (Neuman et al., 2022a; Sridhar, 2023) suggesting that Fmp27 PM targeting relies on a saturable interactor. Using chemicogenomics profiling (further discussed below), we identified the uncharacterized ORF, YBL086C, as the most highly correlated gene to FMP27, and found that overexpression of YBL086C is sufficient to restore the localization of overexpressed Fmp27-ENVY to cortical sites (Sridhar, 2023). The C-terminal portion (residues 216-466) of Ybl086c was necessary and sufficient for Fmp27 recruitment, as well as for co-immunoprecipitation with both Fmp27 and its paralog, Hob2 (Sridhar, 2023). 94 Recruitment and co-immunoprecipitation were lost when alanine substitutions were made in 15 residues within two highly conserved helices in the C-terminus (residues 304-315 for “helix 1”; residues 326-336 for “helix 2”) (Sridhar, 2023). Altogether these findings suggest that Ybl086c, which we will now call Hobbit interactor 1 (Hoi1), is the saturable Fmp27 PM interactor and likely functions in PM recruitment of both Fmp27 and Hob2. 4.2.3 The uncharacterized protein Ybl086c/Hoi1 recruits Fmp27 to the PM Hoi1-mediated Fmp27 PM recruitment has only been demonstrated in the context of overexpressed and C-terminally tagged Fmp27. To determine the necessity of HOI1 for the recruitment of the endogenously expressed protein, confocal imaging of the brighter Fmp27^NG strain was required. First, we validated that, like Fmp27-ENVY, overexpression of internally tagged Fmp27^NG from a strong ADH1 promoter results in a redistribution across the entire ER (Fig 4.3A-B), confirming the loss of ER-PM localization is not due to tag interference upon overexpression. The change in Fmp27 distribution was quantified in an unbiased manner by generating nuclear and cortical ER masks with the Sec63-mScarletI marker and assessing the relative Fmp27^NG intensity levels at each to produce a PM to nuclear ER ratio for quantified cells. On overexpression, the average PM/nuclear ER ratio of Fmp27^NG decreased from 1.6 to 0.8, supporting a redistribution across the ER. When HOI1 was co-expressed from the same strong promoter, ADH1-expressed Fmp27^NG instead showed an enhanced localization to cortical sites at the PM (PM/nuclear ER ratio of 2.0) supporting a role for Hoi1 in Fmp27 recruitment. Moreover, when both Fmp27 and Hoi1 were tagged and overexpressed, both proteins localized to cortical patches with near-perfect colocalization (Fig 4.3C). Altogether, this provides additional support that Hoi1 is a key limiting factor for Fmp27 PM recruitment. 95 Figure 4.3: Uncharacterized protein Hoi1 is the Fmp27 interactor at the PM. (A) Overexpression of HOI1 restores the ER-PM localization of overexpressed (OE) Fmp27^NeonGreen (NG). Confocal images of yeast cells expressing Fmp27^NG and Sec63-mScarletI from their genomic loci. Bar = 2µm. (B) Quantitation of the PM to nuclear ER intensity ratio of Fmp27^NeonGreen (NG) from yeast strains in A determined with line scans. The average intensity ratio is presented. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; cells/replicate/strain = 10; **** = p<0.0001. Error bars indicate SEM. (C) Overexpressed (OE) Fmp27^NeonGreen (NG) colocalizes with overexpressed Hoi1-mScarletI at cortical sites. Fluorescent images of yeast with a linearized line scan and normalized fluorescent signal intensities shown below to highlight the colocalization. The curved arrow in the merged image indicates the line scan start site. Bar = 2µm. As endogenous Fmp27^NG localizes to ER-PM contacts, we wanted to determine if HOI1 is needed for the PM recruitment of endogenously expressed Fmp27. Indeed, in hoi1Δ mutants Fmp27^NG redistributed along the entire ER instead of localizing to cortical puncta (PM/nuclear ER ratio of 0.7) (Fig 4.4A-B). This supports the notion that Hoi1 is necessary for Fmp27 localization to PM contacts. We next imaged a chromosomal copy of Hoi1-ENVY expressed from its native promoter and found that it also localized to cortical puncta (PM/nuclear ER ratio of 1.3) (Fig 4.4C). Unexpectedly in fmp27Δ mutants, Hoi1-ENVY lost its PM localization and became cytosolic (Fig 4.4C-D). Western blotting confirmed that there were no 96 Figure 4.4: The PM localization of Fmp27 and Hoi1 is interdependent. (A) Confocal images of wild-type or hoi1Δ yeast cells expressing Fmp27^NeonGreen and SEC63-mScarletI from their endogenous genomic loci. Bar = 2µm. (B) Quantitation of the PM to nuclear ER intensity ratio of Fmp27^NeonGreen (NG) from yeast strains in A determined with line scans. The average intensity ratio is presented. Individual data points are overlaid and coloured by replicate. Non-parametric two-way t-test; n = 3; cells/replicate/strain = 10; **** = p<0.0001. Error bars indicate SEM. (C) Confocal images of wild-type or fmp27Δ yeast cells expressing Hoi1-ENVY and SEC63-mScarletI from their endogenous genomic loci. Bar = 2µm. (D) Quantitation of the PM to nuclear ER intensity ratio of Hoi1-ENVY from yeast strains in C determined with line scans. The average intensity ratio presented. Individual data points are overlaid and coloured by replicate. Non-parametric two-way t-test; n = 3; cells/replicate/strain = 10; **** = p<0.0001. Error bars indicate SEM. (E) Fmp27-ENVY stability is unaffected by deletion of HOI1. Anti-GFP western blot of Fmp27-ENVY in wild-type and hoi1Δ yeast strains. Pgk1 serves as a loading control. (F) Quantitation of Fmp27-ENVY band intensities in E by densitometry. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; **** = p<0.0001; * = p<0.5; ns = not significant. Error bars indicate SEM. 97 (G) Hoi1-ENVY stability is unaffected by deletion of FMP27. Anti-GFP western blot of Hoi1-ENVY in wild-type and fmp27Δ yeast strains. Pgk1 serves as a loading control. (H) Quantitation of Hoi1-ENVY band intensities in G by densitometry. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; * = p<0.5; ns = not significant. Error bars indicate SEM. (I) Overexpressed (OE) FMP27 restores PM localization of overexpressed Hoi1-mScarletI. Fluorescent images of yeast cells overexpressing Hoi1-mScarletI with endogenous or overexpressed FMP27. Bar = 2µm. (J) Quantitation of the PM to cytosol intensity ratio of Hoi1-mScarletI from yeast strains in I determined with line scans. The average intensity ratio presented. Individual data points are overlaid and coloured by replicate. Non-parametric two-way t-test; n = 3; cells/replicate/strain = 10; **** = p<0.0001. Error bars indicate SEM. changes in Fmp27 or Hoi1 levels when the reciprocal protein was knocked out (Fig 4.4E-H), suggesting Fmp27 is required for Hoi1 PM targeting and that Hoi1 alone is not sufficient for membrane binding. In agreement with this, overexpressed Hoi1-mScarletI was largely cytosolic, although some dim cortical puncta could also be observed (Fig 4.4I-J). Overexpression of FMP27 restored the PM localization of Hoi1-mScarletI confirming that PM recruitment of both proteins is interdependent. In conclusion, we demonstrate that both proteins rely on each other for recruitment to ER-PM contact sites. 4.2.4 The Hoi1 NT-C2 domain is not sufficient for membrane binding Hoi1 has an annotated N-terminal C2 (NT-C2) domain that is predicted to mediate lipid-binding at membranes (Zhang and Aravind, 2010) and a long relatively unstructured C-terminus that mediates the recruitment of Fmp27, and likely Hob2, to PM contact sites (Fig 4.5A-C). As loss of FMP27 results in a loss of membrane localization, this suggests that neither the NT-C2 domain nor C-terminus are sufficient for PM binding. Indeed, overexpressed N-terminal Hoi1- ENVY was unable to localize to membranes in WT, or in FMP27 overexpressing yeast cells (Fig 98 Figure 4.5: The Hoi1 C-terminus is recruited to the ER by Fmp27. (A) Domain schematic of Hoi1/Ybl086c highlighting the annotated NT-C2 domain (PF10358) and conserved helices necessary for interaction with Fmp27. (B) Predicted structure of Hoi1 extracted from the AlphaFold Protein Structure Database (https://alphafold.ebi.ac.uk/; Jumper et al., 2021). The NT-C2 domain and conserved helices are labelled according to the legend in A. (C) The AlphaFold predicted structure of Hoi1 coloured by per-residue pLDDT values. (D) The Hoi1 C-terminus, but not N-terminus, is recruited to the ER by overexpressed (OE) FMP27. Fluorescent images of yeast cells expressing ADH1pr-HOI1-ENVY truncations from plasmids in WT or OE FMP27 strains. Bar = 2µm. 4.5D). In contrast, overexpressed C-terminal Hoi1-ENVY was recruited to the ER in ADH1- FMP27 cells, where the bulk of Fmp27 is located when overexpressed. This demonstrates that the Hoi1 C-terminus is reciprocally recruited by Fmp27 and provides additional support for their interaction. As Hoi1 is necessary for Fmp27 recruitment to PM contacts, but is not sufficient to localize to membranes, we considered the possibility that weak interactions with both PM lipids and PM-localized Fmp27 contribute to targeting of the NT-C2 domain. To test this, Fmp27 must be artificially recruited to the PM. Artificial recruitment was achieved with chimeric constructs where the poorly targeted NT-C2 domain of Hoi1 was replaced with the RFP-labelled PM- 99 Figure 4.6: The Hoi1 C-terminus is sufficient for Fmp27 recruitment but the NT-C2 domain alone is not sufficient for membrane targeting. (A) Schematic of Fmp27 chimeric recruitment assay. The putative membrane-binding NT-C2 domain of Hoi1 is replaced by the PM marker, 2xPH(PLCδ), for enhanced recruitment to PM contact sites. (C) The Hoi1 C-terminus is sufficient for Fmp27 recruitment. Fluorescent images of ADH1-FMP27-ENVY hoi1Δ yeast cells expressing mCherry-2xPH(PLCδ) chimeric constructs from plasmids. Bar = 2µm. (D) Overexpressed (OE) FMP27 artificially recruited to the PM is unable to localize the Hoi1 N-terminus. Fluorescent images of ADH1-FMP27 hoi1Δ yeast cells with plasmids expressing the mCherry-2xPH(PLCδ)-Hoi1(216-466) chimera and Hoi1-ENVY truncations. Bar = 2µm. Schematics of the recruitment assay are shown to the right. localized reporter, mCherry-2xPH(PLCδ), which binds PM-enriched pools of PI(4,5)P2 (Fig 4.6A) (Stefan et al., 2002). When either full-length Hoi1 or its C-terminus were fused to mCherry-2xPH(PLCδ), the resulting chimeric proteins recruited overexpressed Fmp27-ENVY away from the nuclear ER back to cortical patches in cells lacking endogenous HOI1 (Fig 4.6B). However, the plasmid-expressed Hoi1 NT-C2 domain was still unable to target membranes in cells where overexpressed untagged Fmp27 was artificially recruited to the PM by the Hoi1 C-terminus chimera (Fig 4.6C). In contrast, the C-terminus was recruited to both the PM and ER. It is 100 possible that interaction with Fmp27 results in conformational changes in full-length Hoi1 that are necessary for PM targeting of the NT-C2 domain, or that additional interactors may be required for targeting. Alternatively, the conformation of the artificially recruited Fmp27 may not be suitable for NT-C2 domain binding in this context. It is currently unclear how the Fmp27-Hoi1 complex cooperatively targets the PM and further work will be required to determine if this involves interactions with membrane lipids or other contact site proteins. Taken together, we have shown that the C-terminus of Hoi1 is both necessary and sufficient for Fmp27 recruitment, meanwhile the isolated NT-C2 domain is dispensable for the interaction and unable to localize to membranes on its own. 4.2.5 Recruitment by Hoi1 does not require the Fmp27 C-terminus To determine what region of Fmp27 interacts with Hoi1, we generated a series of Fmp27-ENVY C-terminal truncations (Fig 4.7A) and tested their recruitment to cortical sites using the full length mCherry-2xPH(PLCδ)-Hoi1 chimera (Fig 4.7B-D). The majority of the Fmp27 truncation mutants, which had sections of the hydrophobic channel removed, resulted in the formation of bright aggregates when these mutants were overexpressed (Fig 4.7B-D). Interestingly, overexpression of the Hoi1 chimera blocked aggregation and restored cortical PM localization for C-terminal truncations up to Fmp27 residue 1834. Removal of an additional 71 residues, to create truncation mutants Fmp27(1-1763) and shorter, blocked this recruitment. Fmp27(1-1834) is therefore sufficient for Hoi1 interaction (Fig 4.7E). In contrast, the C-terminal 794 residues, which contain repeating hydrophobic ß-groove and ⍺-helical domains, are dispensable. 101 Figure 4.7: Recruitment by Hoi1 does not require the Fmp27 C-terminus. (A) Schematic of Fmp27 indicating the C-terminal truncations tested for PM recruitment by the mCherry-2xPH(PLCδ)-Hoi1 chimera. 102 Truncations that were identified as sufficient for recruitment are labelled in red. (B) Quantitation of the localization of overexpressed (OE) Fmp27-ENVY truncations in live yeast cells expressing plasmids with mCherry-2xPH(PLCδ) only or fused to Hoi1 determined by hand counting. Representative images for each localization category are shown. Bar = 2µm. n = 3; cells/strain/replicate ≥ 306. Unpaired two-tailed parametric T-tests; **** = p<0.0001; *** = p<0.001; ** = p<0.01; * = p<0.5; ns = not significant. Colour of asterisks corresponds to the significance of each representative localization category. Error bars indicate SEM. (C) Representative fluorescent images of overexpressed (OE) Fmp27-ENVY truncations in yeast cells expressing the mCherry-2xPH(PLCδ) plasmid, or (D) mCherry-2xPH(PLCδ)-Hoi1 plasmid from select strains in B. Bar = 2µm (E) AlphaFold predicted structure of Fmp27 (Jumper et al., 2021) indicating the minimal fragment sufficient for Hoi1 recruitment in magenta. (F) AlphaFold predicted structure of Fmp27(1764-1825) coloured by sequence conservation within metazoan homologs as determined by ConSurf (Ashkenazy et al., 2016). The three conserved aspartic acid residues are labelled. (G) Fluorescent images of overexpressed (OE) Fmp27-ENVY mutants in yeast cells expressing the mCherry-2xPH(PLCδ) plasmid, or (H) mCherry-2xPH(PLCδ-Hoi1 plasmid. D>R = Fmp27D1801R; DDD>AAA = Fmp27D1798A,D1801A, 1804A. Bar = 2µm. The Fmp27 truncation recruitment assay suggests that a critical region for the Hoi1 interaction is present between residues 1763 to 1834. Within the AlphaFold model, this region is predicted to form a short segment of the repeating ß-groove structure with a relatively unstructured ~20 residue loop containing three highly conserved aspartic acids (Fig 4.7F). To determine if this loop is necessary for Hoi1-mediated recruitment, we generated Fmp27 mutants with either a substitution of one conserved aspartic acid to arginine (Fmp27D1801R) or substitution of all three aspartic acids to alanine (Fmp27D1798A, D1801A, D1804A). Both mutants appeared to aggregate and mislocalize when overexpressed (Fig 4.7G), and neither were clearly recruited by the mCherry-2xPH(PLCδ)-Hoi1 chimera (Fig 4.7H). While the failure of both Fmp27D1801R and Fmp27D1798A, D1801A, D1804A mutants to be recruited by Hoi1 could result from loss of the Hoi1 binding interface, mutations in this loop could also cause misfolding that impacts localization. Collectively, these results demonstrate that the C-terminus of Fmp27 is dispensable for Hoi1 103 recruitment and suggests the Fmp27 unstructured sequence at residues 1791-1811 is important either for the Hoi1 interaction or for stability. 4.2.6 ColabFold identifies key interacting residues in Fmp27 To identify the residues involved in the Fmp27-Hoi1 interaction we used ColabFold, an AlphaFold2-based protein prediction software (Mirdita et al., 2022) to model the interaction of the Hoi1 C-terminus with Fmp27. Assessing several models generated using different Fmp27 sections from residues 1-1834 identified a predicted protein-protein interaction interface in models using Fmp27(1037-1834) (Fig 4.8A-D). The confidence of these Hoi1(216-466) and Fmp27(1037-1834) models were determined by evaluating the per-residue predicted Local Distance Difference Test (pLDDT) confidence scores. The pLDDT values were largely consistent for Fmp27 across all 5 models, but more variable for Hoi1 with ranked models 1 to 3 providing interpretable values above the confidence threshold (pLDDT value ≥70) (Fig 4.9A). The previously identified conserved Hoi1 “helix 2” and its flanking sequences, including a C-terminal “helix 3”, confidently modeled in contact with Fmp27 at two extended a-helices that project away from the lipid transfer channel (Hoi1 pLDDT scores ≥87) (Fig 4.8B-D, Fig 4.9A). This interaction was consistent among the top 3 models, where the predicted aligned error (PAE) values between interacting residues in Fmp27 relative to interacting residues in Hoi1 were low (< 5 Å) (Fig 4.9B-F), providing additional confidence in the predicted interface. In contrast, no confident interactions were predicted for “helix 1” of Hoi1 (Hoi1 pLDDT scores of ≤65). In the ColabFold models, Fmp27 is predicted to contact the confidently modeled helix 2-3 portion of Hoi1 with 11 residues within its two extended a-helices (Fmp27 pLDDT scores ≥ 104 Figure 4.8: ColabFold identifies key interacting residues in Fmp27. (A) AlphaFold predicted structure of Fmp27 (Jumper et al., 2021) indicating the region used as the ColabFold query in magenta. (B) ColabFold-predicted interaction (Mirdita et al., 2022) between Fmp27(1037-1834) and Hoi1(216-466). The rank 1 model is shown, with residues with pLDDT values <50 hidden. Fmp27 is coloured by per-residue pLDDT value. Hoi1 helices are shown and labelled in gray. (C) ColabFold-predicted interaction between Fmp27(1037-1834) and Hoi1(216-466) as shown in B. Fmp27 is coloured in gray. Hoi1 is coloured by per-residue pLDDT value with helices labelled with green text. (D) A closer look at the ColabFold predicted interface between Fmp27 and Hoi1, in the region indicated by a dashed box in B and C. Fmp27 is coloured in gray and Hoi1 is coloured by per-residue pLDDT values. Side chains are shown for Fmp27 residues predicted to make contact in all 3 top ranked models with pLDDT values >70 for each interacting pair. The identified Fmp27 interacting residues are listed below. (E) Fluorescent images of overexpressed (OE) Fmp27-ENVY or the Fmp271066-1235Δ-ENVY mutant in yeast cells expressing the mCherry-2xPH(PLCδ) plasmids, or (F) mCherry-2xPH(PLCδ-Hoi1 plasmids. Bar = 2µm. (G) Fluorescent images of overexpressed (OE) Fmp27-ENVY or the Fmp2711A-ENVY mutant in 105 yeast cells expressing the mCherry-2xPH(PLCδ) plasmids, or (H) mCherry-2xPH(PLCδ-Hoi1 plasmids. The residues mutated to alanine are those listed in D. Bar = 2µm. 82) (Fig 4.8D). To determine if this region is necessary for Hoi1-mediated PM recruitment, we generated a Fmp271066-1235Δ mutant where the a-helical extension was removed. However, loss of this region resulted in mislocalization of overexpressed Fmp27 to aggregate structures near the cell cortex making the extent of recruitment by the mCherry-2xPH(PLCδ)-Hoi1 chimera uninterpretable (Fig 4.8E-F). As the loss of this region may result in global misfolding, we attempted to generate a milder mutant with alanine substitutions in all 11 identified Hoi1-interacting residues. This Fmp2711A mutant also mislocalized and formed aggregates on overexpression and was not clearly recruited by the Hoi1 chimera (Fig 4.8G-H). It is not apparent if the lack of recruitment by this mutant is due to abolishment of the Hoi1 interface, or due to general misfolding and aggregation. To alternatively dissect this interface and determine the relative contributions of the necessary Hoi1 helices for interaction, we assessed the recruitment of Fmp27 by WT Hoi1, Hoi1helix 1Δ or Hoi1helix 2Δ chimeras (Fig 4.10A). We found that the recruitment of Fmp27 to the PM was significantly reduced when either Hoi1helix 1Δ or Hoi1helix 2Δ chimeras were co-expressed in comparison to the WT chimera, confirming a role for both helices in the Fmp27-Hoi1 interaction. Loss of Hoi1 helix 1 resulted in the greatest reduction in Fmp27 PM recruitment supporting an important role for this helix, even though a binding interface could not be mapped. The moderate recruitment by the Hoi1helix 2Δ chimera is likely due to additional Fmp27 interactions with Hoi1 regions flanking helix 2, in addition to possible interactions mediated by helix 1. 106 Figure 4.9: Confidence measures of the Fmp27-Hoi1 ColabFold models. (A) The per-residue pLDDT values for each of the 5 generated ColabFold models for the Fmp27(1037-1834) and Hoi1(216-466) interaction (Mirdita et al., 2022). The amino acid positions on the left correspond to Fmp27 and those on the right correspond to Hoi1. (B) ColabFold-generated predicted aligned error (PAE) plots for the Fmp27(1037-1834) and Hoi1(216-466) interaction for rank 1, (C) rank 2, (D) rank 3, (E) rank 4, and (F) rank 5 models. To determine if the extended a-helices of Fmp27 contribute to the Hoi1 interface, we generated two new milder Fmp27 mutants to assess their recruitment by Hoi1 chimeric constructs. The first mutant, Fmp27KERKE, has charge swapped substitutions at multiple interacting residues and is predicted to disrupt a larger region of the Fmp27-Hoi1 interface at the Fmp27 extended a-helices (Fig 4.10B-C). The Fmp27KERKE mutant, which localized to the ER on overexpression, showed greatly reduced recruitment by the WT Hoi1 chimera (Fig 4.10A,E) supporting an important role for the extended Fmp27 helices in the Hoi1 interaction. Recruitment of the Fmp27KERKE mutant by either Hoi1helix 1Δ or Hoi1helix 2Δ chimeras further reduced its levels at the PM, with only minimal levels of recruitment observed by the Hoi1helix 2Δ chimera likely because of residual interactions mediated by helix 1. The second mutant, Fmp27NSKR, which has alanine substitutions in 4 interacting Fmp27 residues, is predicted to disrupt interactions with Hoi1 at regions flanking helix 2 and at helix 3 107 Figure 4.10: The Fmp27 ⍺-helical extension interacts with Hoi1. (A) Quantitation of the localization of overexpressed (OE) WT or mutant Fmp27-ENVY in live yeast cells expressing plasmids with mCherry-2xPH(PLCδ)-Hoi1 chimeras as determined by hand counting. Representative images for each localization 108 category are shown to the right. Bar = 2µm. n = 3; cells/strain/replicate ≥ 215 (mean 602). One-way ANOVA with Dunnett’s corrected post-hoc test comparing to WT Fmp27 recruited by the WT Hoi1 chimera; **** = p<0.0001; *** = p<0.001; * = p<0.5; ns = not significant. Colour of asterisks corresponds to the significance of each representative localization category. Error bars indicate SEM. The Hoi1 residues removed in the helix 1Δ chimera are 304-315 and the residues removed in helix 2Δ are 326-336. (B) The ColabFold-predicted interface between Fmp27 and Hoi1. Fmp27 is coloured by per-residue pLDDT values and Hoi1 is coloured in gray. The Fmp27 side chains are shown for residues selected for mutation. (C) Schematic indicating the interacting Fmp27 residues substituted for alanine in the Fmp27NSKR mutant. (D) Schematic indicating the interacting Fmp27 residues substituted for lysine or aspartic acid in the Fmp27KERKE mutant. (E) Fluorescent images of the overexpressed (OE) Fmp27NSKR-ENVY mutant in yeast cells expressing mCherry-2xPH(PLCδ)-Hoi1 chimeras. Bar = 2µm. (F) Fluorescent images of the overexpressed (OE) Fmp27KERKE-ENVY mutant in yeast cells expressing mCherry-2xPH(PLCδ)-Hoi1 chimeras. Bar = 2µm. (Fig 4.10B,D). The Fmp27NSKR mutant localized to the ER on overexpression, and was recruited to the PM by the WT Hoi1 chimera similar to WT Fmp27 (Fig 4.10A,F). However, neither Hoi1helix 1Δ or Hoi1helix 2Δ chimeras were able to recruit Fmp27NSKR. This supports a model where extensive contacts are formed between the Fmp27 ⍺-helical extension and the Hoi1 C-terminus, and suggests that loss of a greater interface is needed to disrupt PM recruitment. Taken together, this mutational analysis demonstrates an important role for the a-helical extension of Fmp27 in Hoi1-mediated PM recruitment. Further, this data supports the hypothesis that both conserved helices within the Hoi1 C-terminus contribute to the Fmp27 interaction, although they may interact at distinct sites. 4.2.7 Fmp27 and Hoi1 have a shared role in lipid homeostasis What is the function of the Fmp27-Hoi1 complex at ER-PM contact sites? To identify functionally correlated proteins that may work together in shared pathways, we took advantage of the homozygous deletion profiling (HOP) chemogenomic dataset to identify yeast genes with 109 Figure 4.11: Fmp27 and Hoi1 are functionally correlated to ER-PM sterol transporters. (A) Ranked genes positively correlated with FMP27 from the homozygous deletion profiling (HOP) chemogenomic dataset (Hoepfner et al., 2014) with p-values <0.01. Genes identified with correlations >0.2 are shown in green with genes under the Lipid Transport GO term (GO:0032365) labelled and shown in dark green. HOI1 (red) is the gene with the highest positive correlation to FMP27. (B) Table of the 10 most highly correlated genes from A with their respective FMP27 correlation values and published localizations listed. (C) Biological processes Gene Ontology (GO) functional enrichment analysis for genes from A with correlations >0.2 (http://geneontology.org). GO terms are presented with their fold enrichment and negative log10 p-values from Bonferroni-corrected Fischer’s exact test. Sterol Transport (GO:0015918), Lipid Transport (GO:0006869) and 110 Intracellular Lipid Transport (GO:0032365) are significantly enriched ontologies. (D) Network of FMP27 and all positively correlated genes from C. Edges indicate the correlation score between nodes with thicker edges indicating higher correlation values. Note the strong correlation between FMP27 and HOI1. Genes are labelled by related functions with retrograde PM-ER ergosterol transporters in green, anterograde ER-PM ergosterol transporters in blue, sphingolipid biosynthetic enzymes in pink, FMP27 in purple and genes with unknown functions in grey. Functionally related genes are denoted with dashed lines. positive correlations to FMP27 (Fig 4.11A-B) (Hoepfner et al., 2014). In this dataset, a diploid library of all non-essential yeast gene deletions was screened on a variety of drugs to determine their relative sensitivities. Using the screen data, we computed correlation values for all genes versus all and extracted a minimal set with significant correlations to FMP27 (p <0.01). Of this subset, the top hits identified with high positive correlations to FMP27 included ER-PM ergosterol transporters and contact site proteins (LAM1/YSP1, LAM2/YSP2/LTC4, LAM3/SIP3, LAF1, OSH3, SCS2) as well as an ER-localized lipid biosynthetic enzyme (ELO2) (Fig 4.11B). Gene Ontology (GO) functional enrichment analysis (http://geneontology.org) of all positively correlated gene using a cut-off of >0.2, identified Sterol Transport (GO:0015918), Lipid Transport (GO:0006869) and Intracellular Lipid Transport (GO:0032365) as significantly enriched ontologies supporting a role for Fmp27 in lipid transport (Fig 4.11C). The most highly correlated gene to FMP27 was HOI1 providing evidence for a shared functional role for both genes. Network analysis of FMP27 and its 10 most correlated genes found similar correlation scores between these genes and both HOI1 and FMP27 (Fig 4.11D), further suggesting that both are functionally linked to sterol transporters at ER-PM contact sites. Furthermore, a drug sensitivity screen of all non-essential haploid deletion mutants identified both FMP27 and HOI1 mutants as some of the most sensitive to the sterol-binding drug, Amphotericin B (AmB) (Parsons et al., 2006; Gray et al., 2012). This anti-fungal drug functions 111 by binding accessible ergosterol, yeast sterol, on the outer leaflet of the PM and mediating membrane permeabilization through channel formation (Gray et al., 2012). Taken together, this suggests that Fmp27 and Hoi1 may have a role in maintaining ergosterol homeostasis, either directly or indirectly. To test sensitivity to AmB, we performed a drug sensitivity spotting assay on a series of mutants in rich yeast peptone dextrose (YPD) media with a sublethal concentration of AmB (1 µM). Knockouts of LAF1 and LAM2 , genes with known AmB sensitivities (Topolska et al., 2020; Gatta et al., 2015) that are highly correlated to both FMP27 and HOI1 (Fig 4.11A,C-D), were used as controls. LAF1 encodes a WD40 repeat-containing protein which interacts with the Lam proteins at ER-PM contacts. LAF1 is required for Lam-mediated retrograde ergosterol transport, and knock-out of this gene, or of the Lam2 transporter, results in accumulation of ergosterol on the PM (Topolska et al., 2020). AmB sensitivity was reproduced in the laf1Δ and lam2Δ mutants and the hoi1Δ mutant was indeed found to be reproducibly sensitive, albeit less sensitive than laf1Δ or lam2Δ (Fig 4.12A). The fmp27Δ and hob2Δ single mutants had no sensitivity, but a double fmp27Δ hob2Δ mutant showed weak sensitivity similar to hoi1Δ suggesting Fmp27 and its paralog Hob2 share some redundant functions. A fmp27Δ hoi1Δ double mutant did not show enhanced sensitivity compared to hoi1Δ suggesting both genes function in the same pathway, while overexpression of both FMP27 and HOI1 resulted in a mild resistance to AmB. Due to the mild sensitivities identified via drug spotting, the AmB sensitivities were further confirmed by a more sensitive plate reader growth assay (Sridhar, 2023). Although sensitivity to AmB suggest increased levels of accessible ergosterol on the outer PM leaflet, sensitivity to this drug can also denote general membrane integrity defects 112 Figure 4.12: Fmp27 and Hoi1 have a shared role in lipid homeostasis. (A) 10-fold serial dilutions of the indicated yeast strains grown on YPD media supplemented with DMSO or 1µM amphotericin B (AmB). Yeast were grown for 3 days at 30°C. (B) Accessible sterols are mislocalized in hoi1Δ mutants. PM localization of the sterol reporter, mCherry-D4H, is rescued with plasmid-expressed wild type or tagged Hoi1, but not by the binding-defective 15A mutant. Fluorescent images of yeast cells expressing TEF1pr-mCherry-D4H from the FF21 locus. Bar = 2µm. (C) Automated quantitation of the average number of internal mCherry-D4H puncta per cell for strains in B. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; cells/strain/replicate ≥ 364; **** = p<0.0001; *** = p<0.001; ** = p<0.01; n.s. = not significant. Error bars indicate SEM. (D) Schematic of the proposed function of Hoi1, Fmp27 and Hob2 in regulating ergosterol distribution. In this model Fmp27 and Hob2, once recruited to the PM by Hoi1, transport sterol-shielding glycerophospholipids from the ER to the PM rendering a pool of ergosterol in the cytosolic PM leaflet inaccessible for transport. Loss of HOI1 or FMP27 and HOB2 thus result 113 in reduced levels of sterol-shielding lipids at PM, increasing the levels of accessible ergosterol that can readily be transported to the ER by Lam2/4 and their regulator, Laf1. (E) Accessible sterols are similarly mislocalized into internal puncta in hoi1Δ and fmp27Δ hob2Δ mutants. Intracellular accumulation of sterols is lost in laf1Δ hoi1Δ double mutants. Fluorescent images of yeast cells expressing TEF1pr-mCherry-D4H from the FF21 locus. Bar = 2µm. (F) Automated quantitation of the average number of internal mCherry-D4H puncta per cell for strains in E. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; cells/strain/replicate ≥ 408; *** = p<0.001; ** = p<0.01. Error bars indicate SEM. (G) Automated quantitation of the average area of mCherry-D4H at the PM per cell for strains in E. Individual data points are overlaid and coloured by replicate. One-way ANOVA with Tukey’s multiple comparison test; n = 3; cells/strain/replicate ≥ 408; *** = p<0.001; ** = p<0.01; * = p<0.5. Error bars indicate SEM. (H) Fluorescent images of yeast cells expressing TEF1pr-mCherry-D4H from the FF21 locus and either plasmid-borne Snc1-GFP or NeonGreen (NG)-ERG6 integrated at the FF18 locus. Blue arrows indicate areas without colocalization; yellow arrowhead indicates colocalization. Bar = 2µm. (Kamiński, 2014; Parsons et al., 2006). As the levels of ergosterol between outer and cytosolic PM leaflets are different (Solanko et al., 2018), we would expect changes in ergosterol homeostasis to modify the distribution at both. To determine if ergosterol distribution within the cytosolic PM leaflet is altered in our HOI1 knock-out mutant, we assessed the distribution of a genetically-encoded fluorescent accessible sterol probe, mCherry-D4H. This probe contains the modified sterol-binding domain 4 (D4) of the C. perfringens theta-toxin with enhanced sensitivity for sterols (Maekawa and Fairn, 2015). When integrated into a safe locus (FF21) and overexpressed using a modified form of the MYLO CRISPR-Cas9 toolkit (Bean et. al., 2022), mCherry-D4H localized to the PM of WT yeast (Fig 4.12B). The distribution was polarized in budding cells, with enhanced D4H signal on the PM of daughter cells. Strikingly, mCherry-D4H was largely absent from the PM in the hoi1Δ mutant and instead accumulated at intracellular puncta (Fig 4.12B-C). This accumulation of mCherry-D4H at internal puncta can be rescued by 114 expressing WT Hoi1 plasmids, but not the Hoi115A mutant with alanine substitutions in conserved residues in helices 1 and 2 that is unable to interact with Fmp27. Is the reduction in accessible ergosterol on the cytosolic PM leaflet the result of a direct loss of sterol transport by Fmp27 and Hob2? The RBG superfamily of bridging LTPs have been shown to preferentially bind and transport glycerophospholipids, but not sterols (Kumar et al., 2018; Valverde et al., 2019; Hanna et al., 2022). We propose a model where Fmp27 and Hob2 transport sterol-shielding lipids, such as phosphatidylserine (PS), to PM contacts (Fig 4.12D). Loss of both paralogs, or of their PM interactor Hoi1, would thus result in a reduction in sterol-shielding lipids on the cytosolic PM leaflet (Maekawa and Fairn, 2015; Kishimoto et al., 2021). Some of these sterols may be flipped to the outer leaflet rendering cells sensitive to AmB, while remaining accessible sterols at the cytosolic leaflet are in turn recognized by the Lam family of LTPs for retrograde transport to the ER. This model could explain why FMP27 and HOI1 show strong functional correlations with the LAM2/4 sterol transporters and their associated regulator, LAF1, and could further explain why the loss of HOI1 causes accessible sterols to accumulate within the cell. To test this hypothesis, we assessed the localization of mCherry-D4H first in fmp27Δ and fmp27Δ hob2Δ mutants (Fig 4.12E-G). As with hoi1Δ, the double fmp27Δ hob2Δ mutant accumulated D4H in intracellular puncta. No significant increase in D4H puncta was observed in the FMP27 knock-out confirming redundancy with HOB2. In a laf1Δ mutant, which is defective in Lam-mediated PM-ER ergosterol transport, a loss of mCherry-D4H polarization between mother and daughter cells was observed with a slight increase in intensity at the PM (Fig 4.12E-G). Notably, when a HOI1 knockout was combined with a LAF1 knockout, the intracellular accumulation of mCherry-D4H was lost and its PM localization restored. The WT sterol 115 distribution in the laf1Δ hoi1Δ double mutant supports our hypothesis that Fmp27 and Hob2 do not directly transport sterols and further suggests that Laf1 acts upstream of HOI1. The site of intracellular sterol accumulation was unclear, but may correspond to the punctate sterol rich compartments (STRICs) identified in the fission yeast S. pombe, which depend on the ER-PM Lam protein, Ltc1, for formation and colocalize with the endosomal marker Syb1 (Marek et al., 2020). Indeed, in hoi1Δ budding yeast occasional colocalization of mCherry-D4H with the Syb1 homolog, Snc1-GFP, was observed but not with another punctate reporter, the lipid droplet marker NG-Erg6 (Fig 4.12H), suggesting accessible sterols may accumulate at related endosomal STRICs in S. cerevisiae. Taken together, we have shown that the Hoi-Fmp27/Hob2 complexes function in sterol homeostasis at ER-PM contact sites in coordination with Laf1 and the Lam transporters. 4.3 Discussion We have determined that the putative MCS-bridging LTP Fmp27 is targeted to PM contacts through interactions with the previously uncharacterized protein Hoi1. Here, we have elucidated the molecular basis of this interaction and find that an extensive interface is formed between a set of conserved ⍺-helices protruding from the Fmp27 hydrophobic channel and the Hoi1 C-terminus. We further demonstrate that complex formation is required for targeting both proteins to ER-PM contact sites. Loss of HOI1 or both FMP27 and HOB2 paralogs results in ergosterol homeostasis defects at the PM supporting a shared function in lipid transport for these complexes, which we demonstrate are in a related pathway with the Lam retrograde sterol transporters. 116 4.3.1 Two conserved Hoi1 helices are required for interaction with Fmp27 We have shown that the extended C-terminus of Hoi1 likely interacts with Fmp27 at two distinct sites. Mutational analysis informed by ColabFold modeling identified an interaction interface between a protruding a-helical extension in Fmp27 and conserved Hoi1 helices 2 and 3. This helical extension observed in Fmp27 is conserved in the paralogous protein Hob2, and is predicted to similarly interact with Hoi1 for PM recruitment. As conserved ⍺-helical extensions characteristically decorate the hydrophobic channels of RBG superfamily LTPs (Neuman et al., 2022b; Levine, 2022), it is interesting to speculate that these regions of other RBG superfamily members could share a role in binding adaptor proteins or regulators. In addition to the ⍺-helical extensions, we identified a distinct conserved C-terminal Fmp27 loop (residues 1791-1811) that is necessary for Hoi1-mediated recruitment but were unable to determine if this site interacted with Hoi1. This Fmp27 loop may directly interact with Hoi1 helix 1, a region that we show is important for Fmp27 recruitment, but which ColabFold does not confidently model. Alternatively, this loop may constitute a binding interface for an additional protein interactor which could bridge an interaction between Fmp27 and Hoi1. This type of 2-step recruitment has been shown for the structurally related Atg2 lipid transporter whose recruitment by its preautophagosomal adaptor, Atg18, relies on additional distinct interactions with Atg9 (Gómez-Sánchez et al., 2018; Chowdhury et al., 2018). An unknown partner may thus bridge a different binding interface between Fmp27 and Hoi1 helix 1, however such a protein would need to be highly abundant as overexpressed Fmp27 and Hoi1 strongly colocalize and co-immunoprecipitate. If Hoi1interacts with Fmp27 at two sites, why is this so? Two interfaces may increase avidity for enhanced recruitment to contact sites. Dual binding sites may also function to orient 117 the Fmp27 channel appropriately at donor and acceptor organelle membranes for efficient lipid transport. Alternatively, two binding sites could be necessary to alter the conformation of Hoi1 to allow for PM association when interacting with Fmp27, possibly by exposing a lipid binding site. Unlike the canonical adaptors for Vps13 (Park et al., 2013, 71; John Peter et al., 2017; Bean et al., 2018), Hoi1 is unable to localize to organelle membranes without Fmp27. We have demonstrated that the Hoi1 NT-C2 domain, which is predicted to bind anionic membrane lipids (Zhang and Aravind, 2010), is necessary for Hoi1 and Fmp27 PM targeting, but is not sufficient. Poor PM affinity of the NT-C2 domain may be enhanced by additional low affinity interactions with the Fmp27 C-terminus when both proteins form a complex. In flies, the last 82 residues of the Fmp27 homolog, Hobbit, are necessary for PM targeting (Neuman et al., 2022a). These residues reside in the annotated APT1 domain, an extension of the lipid transport channel which is sufficient to bind to phosphatidylinositol lipids in vitro (Kolakowski et al., 2020; Kaminska et al., 2016; Neuman et al., 2022a). However, artificially recruiting Fmp27 to the PM did not restore PM localization of the Hoi1 NT-C2 domain. This suggests that conformational changes in full-length Hoi1 or additional protein interactors may be required for PM targeting of this domain. 4.3.2 Role of Fmp27, Hob2 and Hoi1 in ergosterol homeostasis We have determined that loss of both paralogous Fmp27 and Hob2 lipid transporters or their PM recruitment factor, Hoi1, result in altered ergosterol distribution. Unlike WT yeast where the accessible ergosterol reporter mCherry-D4H is localized primarily to the PM, hoi1Δ and fmp27Δ hob2Δ mutants accumulate mCherry-D4H in intracellular puncta. However, this effect on ergosterol distribution does not appear to result from a direct loss of ER-PM ergosterol 118 transport. When deletion of HOI1 is paired with deletion of LAF1, which is necessary for Lam2/Lam4-mediated retrograde PM-ER ergosterol transport (Topolska et al., 2020), mCherry-D4H PM localization is restored. This suggests ergosterol is transported to, but not retained at, the PM in the hoi1Δ mutant and instead is rapidly trafficked away by Lam2/Lam4. What lipids are Fmp27 and Hob2 transporting at the PM? Our HHpred analysis and inspection of AlphaFold predicted structures suggests both paralogs share a similar lipid binding groove to other RBG superfamily members indicating they likely transport related lipid species. Although Vps13 and Atg2 lipid transport channels comigrate with cholesterol in in vitro gel shift assays, both proteins preferentially bind and co-immunoprecipitate with glycerophospholipids (Kumar et al., 2018; Valverde et al., 2019). Recent work on the related yeast protein Csf1 and its C. elegans homolog, LPD-3, also suggest this superfamily member may transport phosphatidylethanolamine (PE) and/or phosphatidylcholine (PC) (John Peter et al., 2022; Toulmay et al., 2022; Wang et al., 2022). Furthermore yeast and human RBG superfamily members have only been demonstrated to transport PE, PC or PS between liposomes (Kumar et al., 2018; Osawa et al., 2019; Valverde et al., 2019; Hanna et al., 2022). Taken together, we hypothesize that Fmp27 and Hob2 similarly transport related glycerophospholipids to the PM indirectly altering ergosterol distribution. The accessibility of ergosterol is modulated by its codistribution with other lipids that can closely associate with and shield ergosterol from proteins such as lipid transporters, biosynthetic enzymes, or synthetic probes (Maekawa and Fairn, 2015; Kishimoto et al., 2021). In yeast, PM ergosterol distribution is asymmetric between bilayers with roughly 80% of PM ergosterol residing within the cytosolic leaflet and 20% in the outer leaflet (Solanko et al., 2018). Sterol-shielding PS and sphingolipids codistribute with sterols in cytosolic and outer leaflets 119 respectively and are required for PM sterol retention (Maekawa and Fairn, 2015; Guan et al., 2009). Reductions in PM PS levels or alterations in the PM phospholipid bilayer distribution of other charged glycerophospholipids, such as PE and PC, reduce total PM sterol levels and cause a relative increase in the proportion of sterols in outer versus cytosolic PM leaflets (Solanko et al., 2018; Kishimoto et al., 2021; Maekawa and Fairn, 2015). Our work identified similar alterations in ergosterol distribution in fmp27Δ hob2Δ or hoi1Δ mutants. The enhanced AmB sensitivity suggests there is a relative increase in ergosterol in the outer leaflet, while redistribution of the cytosolic ergosterol reporter, D4H, is consistent with an overall reduction in sterols within the cytosolic PM leaflet. This supports the hypothesis that Fmp27 and Hob2 transport sterol-shielding glycerophospholipids to the PM and that loss of this transport enhances the accessibility of ergosterol for transport to the ER by Lam2/4. 4.3.3 Conservation of the lipid homeostasis function of Fmp27 and Hoi1 in higher eukaryotes Our work in yeast suggests that Fmp27/Hob2 and Hoi1 function in lipid homeostasis at the PM, possibly in coordination with Laf1 and Lam2/4 ergosterol transporters. Interestingly, in C. elegans the loss of the HOI1 homolog sym-3, or the LAF1 homolog sym-4, causes an enhanced Pharynx ingressed (Pin) phenotype when combined with mutations in fibrillin (fbn-1) or the FBN-1 splicing factor mec-8 (Yochem et al., 2004; Kelley et al., 2015). This Pin phenotype, where the pharynx is detached from the anterior tip of the head, is proposed to result from reduced secretion of apical extracellular matrix (aECM) proteins, in particular FBN-1, which are necessary to resist mechanical forces at the site of pharynx attachment during development (Kelley et al., 2015). As LAF1 is functionally linked to HOI1 in yeast drug 120 sensitivity screens, the shared defects in fbn-1;sym-3 and fbn-1;sym-4 mutants suggest these genes may also function in a shared pathway in nematodes. The FMP27 homolog, F31C3.3, has not been studied and further work is required to determine if F31C3.3 functions with sym-3 and/or sym-4 in nematodes. In flies, the FMP27 homolog hobbit (hob) was originally identified in a screen for mutants with small body size that arrest during metamorphosis (Neuman and Bashirullah, 2018). It was further determined that Hobbit is required for the regulated secretion of insulin-like peptides and mucin proteins, tying members of our complex to secretion in both nematodes and flies. The HOI1 homolog, CG8671, has not been extensively studied but was identified as essential for receptor-mediated endocytosis of dsRNA in a different Drosophila screen (Saleh et al., 2006), and could function as the Hobbit PM adaptor. Are the secretion defects in FMP27 and HOI1 homolog mutants related to PM sterol homeostasis defects? Of note, nematodes are cholesterol auxotrophs and, although they rely on dietary sterol supplementation, sterols are relatively scarce lipids in this organism (Rauthan and Pilon, 2011). Therefore, it is unclear whether the LAF1 and HOI1 nematode homologs regulate the levels of PM sterols, or other lipid species. In yeast, Laf1 and its paralog Dgr2 are b-propeller proteins that regulate the Lam transporters (Topolska et al., 2020). In response to PM perturbing stresses, Lam2/Lam4 are phosphorylated resulting in Laf1/Dgr2 dissociation and inhibition of sterol transport (Roelants et al., 2018; Topolska et al., 2020). It is possible that Laf1/Dgr2 interact with other lipid transporters under these conditions, such as our Fmp27-Hoi1 complex, and could regulate the transport of lipids other than sterols in response to different stresses. PM lipid composition plays a critical role in secretion as lipid microdomains are essential for secretory granule docking, and lipids such as cholesterol, phosphoinositides and 121 glycerophospholipids are necessary for vesicle priming and fusion at the PM (Ammar et al., 2013; Tanguy et al., 2016; Chasserot-Golaz et al., 2010; Zhang et al., 2009; Yang et al., 2021). Of particular interest, cholesterol defines exocytosis sites at the PM and its depletion significantly reduces secretory activity in neuroendocrine cells (Churchward et al., 2005; Chamberlain et al., 2001; Latham et al., 2006) suggesting alterations in sterol homeostasis may also be present in fly hobbit mutants. Other phosphoinositides such as PI(4,5)P2 and PI(3)P positively modulate secretion with enhanced secretory activity observed when PM levels of these lipids are increased, or reduced activity when levels are decreased (Milosevic, 2005; Wen et al., 2008; Meunier et al., 2005). Indeed, fly hobbit mutant cells have altered PI(4,5)P2 distribution with an increase in intracellular PI(4,5)P2 compared to WT (Neuman et al., 2022a), and thus depletion of PI(4,5)P2 from the PM may contribute to the secretory defects. Humans have a single conserved FMP27/HOB2 ortholog known as BLTP2/KIAA0100. BLTP2 was originally identified as overexpressed in breast cancer tissues where its high expression is associated with reduced survival (Song et al., 2006; Liu et al., 2014; Zhong et al., 2018). Silencing BLTP2 in a breast cancer cell line significantly reduces cancer cell viability and additionally reduces cell aggregation, metastasis, and invasion (Zhong et al., 2018). Reduced cell-cell adhesion in BLTP2 knockdowns may result from disruptions in regulated secretion supporting functional conservation. Indeed exogenously expressing BLTP2 in hobbit mutant flies rescues body size and lethality confirming conservation of function as well as supporting a possible conserved localization to ER-PM contact sites (Neuman and Bashirullah, 2018). Overexpressed BLTP2 has been shown to localize to thread-like structures reminiscent of the ER in HEK293 cells (Zhong et al., 2018), similar to the localization of yeast and fly homologs 122 (Neuman et al., 2022a). BLTP2 PM targeting may occur through conserved interactions with HOI1 homologs as the ColabFold-predicted binding interface is conserved. Humans have two HOI1 homologs, EEIG1/FAM102A and EEIG2/FAM102B, with differential expression patterns. RANK-interacting EEIG1 is upregulated by RANK ligand stimulation in bone marrow-derived macrophages undergoing osteoclastogenesis, whereas EEIG2 levels are unaltered (Choi et al., 2013; Jeong et al., 2020). EEIG1 is also upregulated by estrogen supplementation in breast tissue cells (Wang et al., 2004). Differential expression in response to diverse signaling events may function as a switch to activate BLTP2 lipid transport at specific contacts by tethering the C-terminus of the channel to donor membranes when regulated transport is required. This type of regulated recruitment is likely true for the yeast homologs as well, as HOI1 has been identified as upregulated when yeast are supplemented with the phospholipid precursors inositol and choline (Jesch et al., 2005) and down-regulated during anaerobiosis in genome-wide screens (Tai et al., 2005). Further studies will be required to determine which lipids are transported by this conserved complex and to better understand the regulation of lipid transport by Fmp27 homologs and their potential role in secretion. 4.4 Materials and methods 4.4.1 Yeast strains and plasmids Yeast strains and plasmids used in this study are described in Tables 4.1 and 4.2. Plasmids used in this study were made by homologous recombination in yeast by co-transforming linearized plasmids with generated PCR products. Plasmids were recovered in Escherichia coli and sequenced. Yeast strains were similarly made by homologous recombination-based integration of PCR products (Janke et al., 2004; Longtine et al., 1998) 123 unless otherwise stated. Gene deletions and promoter exchanges were confirmed by colony PCR. C-terminal fluorescent tags were confirmed by microscopy or western blot where possible, while tags on low expression strains were confirmed by colony PCR. The Markerless Yeast Localization and Overexpression (MYLO) CRISPR-Cas9 toolkit (Bean et al., 2022) was used to generate strains with stable integration of RPL18B-ymNeonGreen-2xPH(PLCδ). Briefly, yeast were co-transformed with two linearized plasmids; the first providing donor DNA for repair and the second providing Cas9 and gRNA targeting the designated genomic locus. Upon transformation, the Cas9-gRNA plasmid is repaired by homologous recombination at a split-selection marker resulting in restoration of the selectable marker for enhanced efficiency. For stable integration of TEF1-mCherry-D4H, a MYLO toolkit Cas9-gRNA plasmid was co-transformed with a custom-made linearized donor DNA plasmid. Genomic integrations using the MYLO toolkit were confirmed by microscopy. Related CRISPR-Cas9-based genome-editing approaches were used to generate seamless internally ymNeonGreen-tagged FMP27 strains and integrated FMP27 point mutants. Custom linearized Cas9-gRNA plasmids targeting FMP27 sequences were co-transformed with PCR amplified donor DNA. Integration of the internal tag or integration of point mutations was confirmed by sequencing. All validated CRISPR-Cas9 genomically edited strains were grown without selection to lose gRNA plasmids and restore auxotrophy. Table 4.1: Yeast strains used in Chapter 4 Strain ID Genotype BY4741 BY4741 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 VSY007 BY4741 MATa FMP27-ENVY::HIS3 ybl086cΔ::kanMX4 leu2Δ0 met15Δ0 ura3Δ0 VSY005 BY4741 MATa FMP27-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 VSY006 BY4741 MATa YBL086C-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 124 Strain ID Genotype VSY071 BY4741 MATa natMX4::ADH-FMP27-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY683 BY4741 MATa SEC63-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY712 BY4741 MATa YBL086C-ENVY::HIS3 fmp27Δ::kanMX4 leu2Δ0 met15Δ0 ura3Δ0 MDY1321 BY4741 MATa kanMX4::ADH1-FMP27 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 MDY1329 BY4741 MATa natMX4::ADH1-YBL086C-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 MDY1330 BY4741 MATa kanMX4::ADH1-FMP27 natMX4::ADH1-YBL086C-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY884 BY4741 MATa natMX4::ADH1-FMP27-ENVY(1-1911)::HIS3 leu2Δ0 met15Δ0 ura3Δ0 VSY155 BY4741 MATa natMX4::ADH-FMP27(1-996)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 VSY156 BY4741 MATa natMX4::ADH-FMP27(1-1296)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY887 BY4741 MATa natMX4::ADH1-FMP27-ENVY(1-2358)::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY827 BY4741 MATa FMP27^737ymNeonGreen TCB3-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY828 BY4741 MATa FMP27^737ymNeonGreen SEC63-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY831 BY4741 MATa kanMX4::ADH1-FMP27^737ymNeonGreen natMX4::ADH1-YBL086C his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY832 BY4741 MATa kanMX4::ADH1-FMP27^737ymNeonGreen natMX4::ADH1-YBL086C-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 VSY183 BY4741 MATa natMX4::ADH-FMP27(1-1483)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY886 BY4741 MATa natMX4::ADH1-FMP27-ENVY(1-2229)::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY841 BY4741 MATa FMP27^737ymNeonGreen ybl086c∆::kanMX4 SEC63-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY842 BY4741 MATa YBL086C-ENVY::HIS3 SEC63-mScarletI::URA3 leu2Δ0 met15Δ0 SDY843 BY4741 MATa YBL086C-ENVY::HIS3 fmp27Δ::kanMX4 SEC63-mScarletI::URA3 leu2Δ0 met15Δ0 SDY845+ BY4742 MAT⍺ SEC63-mScarletI::URA3 his3Δ200 leu2Δ3 lys2Δ801 SDY846+ BY4742 MAT⍺ sfGFP-SCS2 SEC63-mScarletI::URA3 his3Δ200 leu2Δ3 lys2Δ801 SDY847+ BY4742 MAT⍺ TCB3-EGFP::hisMX6 SEC63-mScarletI::URA3 leu2Δ3 lys2Δ801 SDY848+ BY4742 MAT⍺ FMP27-ENVY::HIS3 SEC63-mScarletI::URA3 leu2Δ3 lys2Δ801 SDY849+ BY4742 MAT⍺ rtn1Δ::kanMX6 yop1Δ::hphNT1 SEC63-mScarletI::URA3 his3Δ200 leu2Δ3 lys2Δ801 SDY850+ BY4742 MAT⍺ rtn1Δ::kanMX6 yop1Δ::hphNT1 sfGFP-SCS2 SEC63-mScarletI::URA3 his3Δ200 leu2Δ3 lys2Δ801 125 Strain ID Genotype SDY851+ BY4742 MAT⍺ rtn1Δ::kanMX6 yop1Δ::hphNT1 TCB3-EGFP::hisMX6 SEC63-mScarletI::URA3 leu2Δ3 lys2Δ801 SDY852+ BY4742 MAT⍺ rtn1Δ::kanMX6 yop1Δ::hphNT1 FMP27-ENVY::HIS3 SEC63-mScarletI::URA3 leu2Δ3 lys2Δ801 VSY195 BY4741 MATa natMX4::ADH-FMP27(1-1381)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 VSY196 BY4741 MATa natMX4::ADH-FMP27(1-1630)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 VSY197 BY4741 MATa natMX4::ADH1-FMP27-ENVY(1-1763)::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY885 BY4741 MATa natMX4::ADH1-FMP27-ENVY(1-1973)::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY921 BY4741 MATa natMX4::ADH1-FMP27(1-1834)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 SDY981 BY4741 MATa kanMX4::ADH1-FMP27 ybl086c∆::natMX4 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1002 BY4741 MATa ybl086c∆::kanMX4 FF21::TEF1-mCherry-D4H his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1011 BY4741 MATa kanMX4::ADH1-FMP27^737ymNeonGreen SEC63-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY1012 BY4741 MATa kanMX4::ADH1-FMP27^737ymNeonGreen natMX4::ADH1-YBL086C SEC63-mScarletI::URA3 his3Δ1 leu2Δ0 met15Δ0 SDY1015 BY4741 MATa ybl086cΔ::kanMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1016 BY4741 MATa fmp27∆::kanMX4 ypr117wΔ::natMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1017 BY4741 MATa fmp27Δ::kanMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1020 BY4741 MATa ybl086c∆::kanMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-ERG6-NeonGreen his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1022 BY4741 MATa FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1026 BY4741 MATa laf1Δ::kanMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1027 BY4741 MATa kanMX4::ADH-FMP27 natMX4::ADH-YBL086C FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 SDY1031 BY4741 MATa laf1∆::kanMX4 ybl086c∆::natMX4 FF21::TEF1-mCherry-D4H FF18::RPL18B-NeonGreen-2xPH(PLCδ) his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 126 Strain ID Genotype MDY1393 BY4741 MATa natMX4::ADH1-FMP27(W1796A,D1798A,D1801A,F1802A,D1804A)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 MDY1396 BY4741 MATa natMX4::ADH-FMP27(N1185A,S1189A,K1192A,R1193A)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 MDY1409 BY4741 MATa natMX4::ADH-FMP27(D1798A,D1801A,D1804A)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 MDY1410 BY4741 MATa natMX4::ADH-FMP27(D1801R)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 MDY1425 BY4741 MATa natMX4::ADH-FMP27(K1090D,E1093K,R1097D,K1192D,E1196K)-ENVY::HIS3 leu2Δ0 met15Δ0 ura3Δ0 + Parental strains received from Dr. W. Kukulski (University of Bern). Table 4.2: Plasmids used in Chapter 4 Plasmid ID Plasmid Description Source pRS413 pRS413 (HIS3, CEN) This study pCS07 pRS316-UTR-SNC1-GFP (URA3, CEN) This study pRS415 pRS415 (LEU2, CEN) This study pRS413- YBL086C pRS413-UTR-YBL086C (HIS3, CEN) This study pVS002 pRS413-UTR-YBL086C-ENVY (HIS3, CEN) This study pVS006 pRS413-UTR-YBL086C(216-466)-ENVY (HIS3, CEN) This study pVS008 pRS413-UTR-YBL086C(1-216)-ENVY (HIS3, CEN) This study pYL95 pRS415-GDP1-mCherry-2xPH(PLCδ) (LEU2, CEN) (Omnus et al., 2020) pVS019 pRS415-GDP1-mCherry-2xPH(PLCδ)-YBL086C (LEU2, CEN) This study pVS020 pRS415-GDP1-mCherry-2xPH(PLCδ)-YBL086C(216-466) (LEU2, CEN) This study pVS036 pRS415-GDP1-mCherry-2xPH(PLCδ)-YBL086C(304-315Δ) (LEU2, CEN) This study pVS037 pRS415-GDP1-mCherry-2xPH(PLCδ)-YBL086C(326-336Δ) (LEU2, CEN) This study pVS044 pRS413-UTR-YBL086C(M302A,L305A,S308A,L309A,Y310A,T313A, F314A, L316A,V331A,E332A,D333A,I334A,L335A,G337A,G338A)-ENVY (HIS3, CEN) This study 127 4.4.2 Microscopy image acquisition and processing Yeast cells were cultured overnight in synthetic minimal media (SD) (0.68% [w/v] yeast nitrogen base, 2% [w/v] ammonium sulfate, amino acid supplements) at 30 °C, then diluted and grown to logarithmic phase. Cells were transferred to 96-well glass-bottom plates (Eppendorf or Cellvis) pre-treated with concanavalin A (Sigma-Aldrich). After 4 minutes, wells were aspirated to remove non-adherent cells and fresh SD media was added back for imaging. Microscopy images were acquired at room temperature on a Leica TCS SP8 Super-Resolution Confocal Microscope (Leica Microsystems, Wetzlar, Germany) with a high-contrast Plan Apochromat 63×/1.30 Glyc CORR CS objective (Leica Microsystems, Wetzlar, Germany) and an ORCA-Flash4.0 digital camera (Hamamatsu Photonics, Shizuoka, Japan). Fluorescent and DIC images were acquired and processed with MetaMorph 7.8 software (MDS Analytical Technologies, Sunnyvale, California). For presentation, images were cropped in Photoshop CC 2022 (Adobe, Mountain View, California) and the brightness and contrast adjusted linearly for the same fluorescent proteins so that quantitative differences in fluorescent intensity are represented. Confocal images were acquired with Leica Application Suite X (LASX) 3.5.7 software (Leica Microsystems, Wetzlar, Germany) (490nm for NeonGreen/ENVY/EGFP/sfGFP and 570nm for mScarletI) and subsequently deconvolved using Huygens Essentials software (Scientific Volume Imaging, Hilversum, Netherlands). Representative deconvolved confocal images were individually adjusted for brightness and contrast in Photoshop CC 2022 (Adobe, Mountain View, California). 128 4.4.3 Quantitation of microscopic data For representative line scans to visualize colocalization, line profiles of the cell cortex were extracted and linearized using Fiji (Schindelin et al., 2012). Fluorescent signal intensities were normalized to absolute values and plotted. To determine PM to nER intensity ratios, cells were identified with clearly separated nuclei and cell cortexes using the ER marker Sec63-mScarletI. For each cell, selections were made around the cell cortex (PM) and the nER in Sec63-mScarletI images and propagated to the corresponding Fmp27^NeonGreen or Ybl086c-ENVY images using Fiji (Schindelin et al., 2012). Fluorescent signal intensities across each pixel were extracted and the average intensity used in computing the PM to nER intensity ratio for each cell. To determine PM to cytosol intensity ratios, selections were made around the cell cortex (PM) and a separate selection made within the cell in DIC channel images and propagated to YBL086C-mScarletI images using Fiji (Schindelin et al., 2012). Fluorescent signal intensities across each pixel were extracted and the average intensity used in computing the PM to cytosol intensity ratio for each cell. Statistical analyses were carried out by performing either non-parametric two-way T-tests for experiments with two strains, or one-way ANOVA with post-hoc Tukey’s multiple comparison test for experiments with three strains using GraphPad Prism 9.1.0 (GraphPad Software; San Diego, California). Data distribution was assumed to be normal, but not formally tested. For automated quantitation of the number of mCherry-D4H internal puncta per cell and mCherry-D4H area at the PM, images were analyzed using a custom MetaMorph 7.8 journal (MDS Analytical Technologies, Sunnyvale, California) as described below. All strains analyzed had the mCherry-D4H accessible sterol reporter and the NeonGreen-2xPH(PLCδ) PM reporter integrated at separate safe loci. Fluorescent images were captured with GFP, RFP and BFP filter 129 sets. Live cells were identified in the BFP channel using the Count Nuclei function. Dead cells and extracellular noise in GFP, RFP and BFP channels were identified with iterative Count Nuclei functions using high intensity thresholds and masked with the Arithmetic function. A final Count Nuclei computation was used on the masked images to generate a live cells mask. The PM was identified with the Morphological Top Hat function in the GFP channel using the NeonGreen-2xPH(PLCδ) marker. Identified PM regions were selected with the Count Nuclei function, inverted, and eroded. A PM masked image was generated by combining the live cells mask and eroded PM image using the Arithmetic function. An inverse masked image of only the PM was similarly generated by combining the live cells mask with an inverted eroded PM mask using the Arithmetic function. mCherry-D4H signals were identified with the Granularity function and combined with the live cells mask to identify only mCherry-D4H signals in live cells. Using the PM masked image, an additional Granularity computation was used to identify only intracellular mCherry-D4H puncta. Intracellular puncta per cell counts were computed by dividing the number of intracellular puncta within an image by the number of live cells identified in that image. mCherry-D4H signals at the PM were identified with the Granularity function on the PM only masked image and the cumulative size of all identified regions within an image computed and divided by the number of live cells identified in that image. Statistical analyses were carried out by performing one-way ANOVAs with post-hoc Tukey’s multiple comparison test using GraphPad Prism 9.1.0 (GraphPad Software; San Diego, California). Data distribution was assumed to be normal, but not formally tested. 130 4.4.4 Western blot analysis of Fmp27-ENVY and Ybl086c-ENVY Yeast cells were cultured overnight in complete yeast peptone dextrose media (YPD) (1% [w/v] yeast extract, 2% [w/v] Bacto-Peptone, 2% [w/v] glucose, 0.032% [w/v] tryptophan) at 30 °C, then diluted and grown to logarithmic phase. 10 OD600/mL equivalent of cells were harvested, washed, and stored at -80 °C for 20 minutes. Cells were resuspended in 100µL Thorner buffer (8M urea, 5% SDS, 40nM Tris-Cl (pH 7.0), 1% b-mercaptoethanol and 0.4 mg/mL bromophenol blue) and lysed by vortexing with 0.5 mm diameter glass beads three times at 45 second intervals with 1 minute incubation on ice between vortexes. Proteins were denatured at 70 °C for 5 minutes. For Fmp27-ENVY, 1 OD600/mL equivalent of cell lysate was separated on 7% SDS-PAGE gels. For Ybl086c-ENVY and endogenous PGK1, 0.4 and 0.5 OD600/mL equivalent of cell lysate respectively was separated on 10% SDS-PAGE gels. Proteins were transferred from SDS-PAGE gels to nitrocellulose membranes (50-206-3328; Fisher Scientific) and blotted with either mouse monoclonal a-GFP (11814460001; Roche) or a-PGK1 (22C5D8; Thermo Fisher Scientific) antibodies, followed by horseradish peroxidase-conjugated polyclonal goat a-mouse (115–035-146; Jackson ImmunoResearch Laboratories) antibodies. Blots were developed with ECL chemiluminescent reagents (RPN2106, Cytiva) and exposed to Amersham Hyperfilm ECL (CA95017-653L; VWR). Developed films were scanned, and densitometry was performed in Fiji (Schindelin et al., 2012). Statistical analysis was carried out by performing a one-way ANOVA with post-hoc Tukey’s multiple comparison test using GraphPad Prism 9.1.0 (GraphPad Software; San Diego, California). Data distribution was assumed to be normal, but not formally tested. 131 4.4.5 Amphotericin B sensitivity spot assay Yeast cells were cultured overnight in complete yeast peptone dextrose media (YPD) (1% [w/v] yeast extract, 2% [w/v] Bacto-Peptone, 2% [w/v] glucose, 0.032% [w/v] tryptophan) at 30 °C, then diluted and grown to logarithmic phase. 1 OD600/mL equivalent of cells were harvested and 10-fold serial dilutions prepared. 2 µL of each dilution was spotted onto complete YPD plates supplemented with DMSO or 1µM amphotericin B (A4888; Sigma-Aldrich). Plates were incubated at 30 °C and imaged after 3 days. 4.4.6 Protein-protein interaction modeling Ab-initio prediction of protein-protein interactions was performed by the ColabFold 1.3.0 AlphaFold2-Multimer MMseq2-driven homology search software (Mirdita et al., 2022; https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb) with default settings. Analysis of predicted contact residues was performed in UCSF ChimeraX version 1.4.0 (Pettersen et al., 2021; University of California San Francisco Parnassus Campus). Residues in contact were identified with the standard atom VDW overlap parameter of ≥0.40 Å. Predicted contact residues were further refined by evaluating predicted aligned error (PAE) domains and pruning contact residues that were found in separate PAE domains. Figures were generated in UCSF Chimera version 1.16.0 (Pettersen et al., 2004; University of California San Francisco Parnassus Campus) and assembled in Adobe Illustrator 2022 (Adobe, Mountain View, California). 132 4.4.7 Chemicogenomic dataset and gene ontology analysis The homozygous deletion profiling (HOP) chemicogenomics dataset (Hoepfner et al., 2014) was analyzed with RStudio (RStudio Team, Boston, Massachusetts). Briefly, the raw screen values were extracted and Pearson correlation coefficients computed for all versus all gene comparisons. Genes with positive genetic correlations to FMP27 and with a p-value <0.01 were extracted to generate a smaller subset that were pruned in Cytoscape version 3.5.1 (Shannon et al., 2003) to generate a network of genes with significant correlations to FMP27 (2833 nodes and 2832 edges). The correlation values were plotted in Microsoft Excel 2022 (Microsoft, Redmond, Washington) while the outputted graph and accompanying table were formatted in Adobe Illustrator 2022 (Adobe, Mountain View, California). The network of FMP27 and positively correlated genes was made in Cytoscape version 3.5.1 (Shannon et al., 2003). Gene Ontology (GO) functional enrichment analysis was performed using the Gene Ontology Consortium (Ashburner et al., 2000; The Gene Ontology Consortium et al., 2021) Enrichment Analysis tool (Mi et al., 2019). The Saccharomyces cerevisiae reference was used and GO biological process complete annotation data set selected for analysis. Fisher’s Exact test and Bonferroni correction for multiple testing was applied. 133 Chapter 5: Discussion and Conclusions 5.1 Summary of Key Findings The studies described here contribute to our understanding of the membrane targeting strategies of Vps13 and related bridging lipid transporters. Understanding how VPS13 proteins target organelle membranes is essential to understanding their functions and roles in disease, as dysregulation of each of the four human homologs is implicated in distinct neurological disorders which are proposed to arise from the loss of lipid transport between specific organelle pairs (Rampoldi et al., 2001; Kolehmainen et al., 2003; Lesage et al., 2016; Seong et al., 2018; Gauthier et al., 2018). Using budding yeast as a model, we determined the necessity of a related motif in all organelle-specific adaptor proteins for the recruitment of the single conserved yeast Vps13. We showed these related motifs compete for binding to a conserved six-repeat region, which we have named the Vps13 Adaptor Binding (VAB) domain, and further identified a minimal interaction site in the latter two repeats. A causative VPS13D spastic ataxia mutation that alters a conserved residue in the minimal adaptor-binding site blocked adaptor interaction and Vps13 recruitment when modeled in yeast. These findings indicate that yeast Vps13 localization is coordinated by interactions with adaptor proteins, which may be further modulated by changes in relative adaptor expression levels and/or by regulated exposure of the adaptor binding motifs. Further, this work suggests that VPS13 VAB domains have a conserved adaptor-binding function in higher eukaryotes. After Vps13 was found to form an elongated lipid transfer channel through structural reconstruction by X-ray crystallography or cryo-EM (Li et al., 2020; Kumar et al., 2018), we used bioinformatic approaches to identify the putative structural homologs, Fmp27/Hob1 and Ypr117w/Hob2. We determined that like Vps13, Fmp27 relies on a previously uncharacterized 134 protein interactor, Ybl086c/Hoi1, for recruitment to ER-PM contact sites. We further identified a large interaction interface between the Hoi1 C-terminus and an ⍺-helical extension of Fmp27, which is conserved in Hob2. Loss of either the PM interactor, or both paralogous lipid transporters, results in a redistribution of accessible ergosterol from the PM to intracellular puncta that requires retrograde sterol transport by the Lam proteins. Altogether, these findings provide evidence for a related lipid transfer function for Fmp27 and Hob2 at MCSs supporting their classification as new members of the RBG superfamily of bridging lipid transporters. 5.2 Vps13 adaptor proteins bind a single site in the VAB domain We have determined that the six-repeat VAB domain of yeast Vps13 interacts with adaptor proteins through related motifs we have named proline-X-proline (PxP) motifs, due to the necessity of these prolines for Vps13 interaction and recruitment. The adaptor PxP motifs bind at VAB repeats 5-6 and mutation of a conserved asparagine residue in repeat 6 is sufficient to disrupt interaction, suggesting a single adaptor binding site lies within this region. Our conclusions for the necessity and sufficiency of these regions for Vps13-adaptor interaction were based on co-immunoprecipitation experiments or fluorescent microscopy recruitment assays. A main limitation for these assays is that they do not demonstrate direct binding and thus cannot rule out that the Vps13-adaptor interaction is bridged by additional protein interactors. After publication of the manuscript described in Chapter 3, the Reinisch research group published the resolved crystal structure of the C. thermophilum Vps13 VAB domain bound to the Mcp1 PxP motif-containing peptide from the same organism, conclusively determining a direct interaction between Vps13 and adaptor proteins (PDB 7U8T; Adlakha et al., 2022). In the resolved structure, each VAB domain repeat forms a ß-sandwich fold, similar to AlphaFold2-135 predicted models. The Mcp1 PxP motif binds the VAB domain primarily at repeat 6 with additional interactions formed by residues in repeat 5 (Adlakha et al., 2022), confirming this region as the yeast adaptor-binding site. In the crystallized proteins the Mcp1 PxP peptide was fused to the N-terminus of the VAB domain and, due to the distance from N- to C-termini, the VAB-adaptor interactions occurred in trans between separate monomers resulting in dimer formation that was detected by size exclusion chromatography. Related constructs with the S. cerevisiae PxP motifs of each adaptor protein (Ypt35, Mcp1 and Spo71) fused to the N-terminus of the budding yeast VAB domain, but not a VAB only construct, similarly dimerized. Dimerization was blocked by mutating homologous interacting residues in the VAB domain, as identified in the C. thermophilum crystal structure (Adlakha et al., 2022), supporting the idea that all 3 adaptors bind the same pocket within the VAB domain. Our mutational analysis of the VAB domain determined that mutation of conserved asparagine residues in repeats 1 and 6 were sufficient to block adaptor interactions. In the resolved structures, the conserved asparagine of each repeat was found at the end of a ß-sheet where it formed hydrogen bonds with other residues within the same repeat, likely stabilizing the ß-sandwich structure (Adlakha et al., 2022). The asparagine in repeat 6 lies near the binding pocket but does not form direct interactions with the PxP peptide, and thus mutation of this residue likely results in local misfolding that disrupts the binding pocket. The reasoning behind the effect on adaptor binding of a similar mutation in repeat 1 is less clear as local misfolding in a distant repeat is unlikely to affect the PxP binding site. In support of this, our data suggests mutations in VAB repeats 2 to 5 do not strongly disrupt adaptor interactions and, in fact, naturally occurring asparagine to serine substitutions are present in these repeats in members of the division Magnioliophyta and Ascomycota, as well as in human VPS13B. Localized 136 misfolding caused by mutation of VAB repeat 1 may instead alter the positioning of the VAB domain in relation to the Vps13 lipid transfer channel or membranes due to its proximity to the channel. Assays that detect protein unfolding, such as differential scanning fluorimetry (DSF) or differential static light scattering (DSLS) (Senisterra et al., 2010; Gao et al., 2020; Senisterra and Finerty, 2009), will be needed to determine if the VAB domain mutants are intrinsically more disordered. In our original publication we had proposed that the VAB domain is comprised of a series of ß-propeller folds due to structural predictions of yeast and human VAB domains generated with the I-TASSER suite (Dziurdzik et al., 2020), which were in line with other predictions at the time that suggested a WD40-like fold (Kumar et al., 2018; Ugur et al., 2020). Later, with the release of AlphaFold2, we updated our model to propose that this domain forms a series of ß-sandwiches instead (Dziurdzik and Conibear, 2021). The discrepancy between these models can be explained by the different approaches used by each prediction tool. I-TASSER relies on first threading the query to resolved protein structures and thus is biased towards known structures in repositories (Yang et al., 2015). AlphaFold instead relies on co-evolution of amino acids, which can indicate proximity in protein structures or functional importance, and first generates multiple sequence alignments with the query to develop constraints that guide structure prediction (Jumper et al., 2021). Our AlphaFold2 predicted VAB domain models were similar to the resolved structure (an overall C⍺ RMSD of 1.135 Å across 305 pruned paired residues). As noted by the Reinisch lab, AlphaFold2-predicted structures accurately predicted and superimposed closely in the first 4 VAB repeats but differed more at the C-terminal repeats (Adlakha et al., 2022). The reduced superimposition of the latter repeats may be an artifact of crystal packing or could arise from the additional constraint resulting from the in trans 137 interaction between different PxP-VAB monomers. The higher accuracy predictions generated by AlphaFold2 in comparison to I-TASSER are not surprising due to the specificity of the VAB domain to VPS13 proteins and lack of published structures at the time of model generation. The high confidence structure predictions made by AlphaFold2 for uncharacterized proteins make it a powerful tool for hypothesis generation. 5.3 Additional VAB-interacting adaptor proteins in yeast and humans Three PxP motif-containing adaptors have been identified that recruit Vps13 to either endosomes/vacuoles, mitochondria or the prospore membrane (John Peter et al., 2017; Park et al., 2013; Bean et al., 2018), and it remains to be determined if additional adaptors are required for recruitment to other organelles where Vps13 has been reported, such as peroxisomes . Our work suggests that another adaptor protein is required for the carboxypeptidase Y (CPY) sorting function of Vps13 as mutations that disrupt adaptor interactions, or overexpression of soluble PxP motifs, result in CPY secretion. To identify additional putative yeast adaptor proteins, high-throughput protein-protein interaction screening methods, such as tandem-affinity purification and mass spectrometry (TAP-MS) or yeast 2-hybrid (Y2H)-based screens, could be performed with the Vps13 VAB domain as bait. TAP-MS provides high specificity for interactors due to two successive affinity purification steps that reduce the number of false positive interactors identified, however this technique is thus worse at identifying weak or transient interactors (Gerace and Moazed, 2015; Adelmant et al., 2019). Additionally, TAP-MS or related immunoprecipitation followed by mass spectrometry (IP-MS) screens only capture the repertoire of putative interactors under the specific physiological conditions and cell state tested (Gerace and Moazed, 2015; Adelmant et 138 al., 2019). Y2H screens, on the other hand, can assess numerous peptide interactions without requiring the specific cell conditions needed for the in vivo protein interaction to occur (Brückner et al., 2009; Mehla et al., 2017), which may be crucial for identifying novel Vps13 adaptors. For example, Spo71 has the strongest recruitment signal of the three known adaptors but is expressed only during sporulation (Park et al., 2013; Bean et al., 2018) and could be missed by TAP-MS if sporulation was not induced. Y2H screens have lower specificity but are thus more likely to identify weak or transient interactors making them better suited for the identification of Vps13 adaptors (Brückner et al., 2009; Koegl and Uetz, 2007), which rely on variable recruitment signal strengths to compete for binding to a single site (Bean et al., 2018). In particular, dynamic enrichment for evaluation of protein networks (DEEPN), a Y2H screening method that relies on competition between different plasmid-bearing populations followed by high-throughput sequencing of the barcoded population and computation, would be well suited for the identification of additional transient interactors (Pashkova et al., 2016). One major limitation for Y2H screens is that they require the import of bait and prey peptides into the yeast nucleus and thus may not detect integral membrane protein interactors (Koegl and Uetz, 2007). The VAB domain may act as a protein scaffold and bind additional proteins at other sites besides repeat 6. Loss of such a binding partner resulting from mutations in VAB repeat 1 could affect Vps13 recruitment to membranes and disrupt PxP adaptor interactions. These interactors could similarly be identified with TAP-MS or DEEPN Y2H screens and be differentiated from canonical adaptors by including a binding defective VAB domain bait as an additional control. For example, a concurrent screen could be performed with a mutant VAB domain containing substitutions in the PxP motif-interacting residues identified from the crystal structure in addition to a screen with the wildtype VAB domain. Comparisons of the relative interactions detected by 139 either wildtype or mutant baits can then be used to differentiate between interactors that bind the adaptor pocket in repeat 6 from those that bind elsewhere in the VAB domain. Our work suggests that human VAB domains have a conserved adaptor-binding function because modeling of the homologous VPS13D N3521S disease-associated substitution in VAB repeat 6 blocked adaptor interactions in yeast. In support of a necessary role for this region, plasmid-expressed VPS13D with the N3521S substitution is unable to rescue mitochondrial morphology defects in VPS13D knockout cells (Baldwin et al., 2021). Further, the VPS13D VAB domain has now been found to interact with mitochondrial Miro, and a peroxisome-enriched splice variant, and these interactions recruit VPS13D to mitochondrial contact sites (Guillén-Samander et al., 2021). Beyond mitochondrial recruitment, the VPS13D VAB has also been shown to interact with TSG101 for recruitment to lipid droplets after oleic acid supplementation suggesting human VAB domains may also possess a larger repertoire of interactors (Wang et al., 2021). In addition to VPS13D, other human VPS13 VAB domains have also been implicated in membrane targeting or adaptor interactions. A VAB domain-containing fragment of VPS13C is sufficient for targeting to endolysosomal membranes (Kumar et al., 2018) and a disease-causing VPS13A homozygous missense mutation in VAB repeat 6 perturbs VPS13A localization to LDs (Park and Neiman, 2020; Velayos-Baeza et al., 2008). The VPS13A VAB has further been demonstrated to interact with SNX5, a sorting nexin at endosomes, with a PxP-like sequence necessary for the interaction (Tornero-Écija et al., 2023). The VAB domains therefore have conserved adaptor binding roles in at least some human VPS13 proteins. However, if the 4 homologs have diverged to localize to different subsets of MCSs they must use different localization determinants. Fine-tuning of adaptor motifs and VAB domain adaptor-binding sites could allow different VPS13 proteins to recognize 140 different adaptors. Indeed, adaptors and adaptor motifs seem to undergo rapid evolutionary changes: the consensus PxP motif used in S. cerevisiae is conserved only within the order Saccharomycetales and homologs of the mitochondrial and prospore membrane adaptors Mcp1 and Spo71 are found only in fungi. Although the VPS13A-SNX5 interaction relies on a related PxP motif (Tornero-Écija et al., 2023), this sequence is located in the phospholipid-binding Phox domain and is involved in intramolecular interactions that play a structural role and thus may not contribute directly to binding (Koharudin et al., 2009). To identify human adaptor proteins and determine the adaptor motifs bound by each homolog, related TAP-MS or DEEPN Y2H screens can be performed with each homolog VAB domain as bait. Here, TAP-MS may be more suitable as a number of post-translational modifications (PTMs) in human proteins are not available in yeast. If such PTMs are required for VAB domain interactions, the relevant interactors would not be identified with a Y2H screen. For example, the VPS13D VAB domain contains a ricin B-type lectin domain that is thought to bind N-acetylgalactosamine (GalNAc), a PTM not found in yeast, which may contribute to adaptor interactions (Dall’Armellina et al., 2023; de Las Rivas et al., 2019). However, as the relevant cell types can be difficult to determine, especially with the ubiquitous expression of human VPS13 proteins, a DEEPN Y2H screen could provide a useful first look. After identification of putative interactors, the hits will need to be validated with subsequent Y2Hs and co-immunoprecipitation experiments. After validation, the interacting adaptor motifs can be modeled with their respective VAB domain using AlphaFold2 Multimer to identify the putative interacting residues and the necessity of these motifs tested experimentally by co-immunoprecitation. 141 5.4 Drivers of lipid transport by bridging channels Vps13 and Atg2 form elongated channels that bridge organelle membranes and allow lipids to flow between them. This mechanism of lipid transfer could provide a rapid supply of bulk lipids to organelles to function in membrane expansion events, such as prospore membrane expansion during sporulation (Vps13) (Park and Neiman, 2012) or autophagosome formation during macroautophagy (Atg2) (Chowdhury et al., 2018; Osawa et al., 2019). Unidirectional transport would be required for such membrane expansion events to occur, however Atg2 has been shown to transport lipids bidirectionally between liposomes in vitro (Maeda et al., 2019). Other interactors or proteins in proximity to Atg2 at ER-phagophore contacts may contribute to directionality as well as the rate of transport. In support of this, the rates of Atg2 lipid transfer determined by in vitro liposome-based transfer assays is slower than that of shuttling lipid transporters and appears too slow to drive autophagosome expansion in vivo (Maeda et al., 2019; Valverde et al., 2019). Other protein interactors for Atg2 and Vps13 may include enzymes that drive lipid loading into the channels through ATP-dependent processes or lipid biosynthetic enzymes that generate a lipid source on acceptor membranes or lipid sink on donor membranes. For example, lipases and phospholipid synthases localize at sites near Atg2 to provide newly synthesized lipids that can be rapidly channeled into Atg2 during autophagy (Schütter et al., 2020; Andrejeva et al., 2019; Dupont et al., 2014; Kannan et al., 2017). One example of a unidirectional lipid transporting bridge is the lipopolysaccharide (LPS) transport (Lpt) complex in gram-negative bacteria that transports LPS from the inner to outer membranes (Reinisch and Prinz, 2021; Wong et al., 2019). This complex relies on ATP-binding cassette transporters to load lipids into their channel and drive unidirectional transport through ATP-hydrolysis (Sperandeo et al., 2017). However, other bridging lipid transporters in gram-142 negative bacteria have been identified with structural homology to Vps13 and Atg2 (TamB/AsmA family) and these have instead been proposed to transfer lipids bidirectionally by diffusion (Grimm et al., 2020; Ruiz et al., 2021; Neuman et al., 2022b), leaving the directionality of transport by Vps13 and related proteins an open question. As Vps13 and Atg2 are proposed to transfer lipids between cytosolic membrane leaflets, this necessitates the requirement for coupled transbilayer lipid transporters to prevent lipid accumulation or depletion from individual leaflets. Indeed, both Vps13 and Atg2 have now been found to interact with scramblases on membranes. Atg2 interacts with the scramblase Atg9 at the isolation membrane/phagophore (Kotani et al., 2018a; Matoba et al., 2020; Maeda et al., 2020) and with VMP1 and TMEM41B at the ER (Ghanbarpour et al., 2021), ensuring membrane bilayers on either end of the lipid transfer channel are equilibrated. For now, Vps13 has only been found to interact with scramblases at its C-terminus. In yeast the mitochondrial adaptor, Mcp1, has been demonstrated to function as a scramblase (Adlakha et al., 2022) and this activity is needed for ERMES rescue by Vps13 (John Peter et al., 2017). Meanwhile human VPS13A interacts with the XK scramblase through its PH domain (Guillén-Samander et al., 2022; Park and Neiman, 2020; Adlakha et al., 2022). Interestingly, mutations in XK are associated with McLeod syndrome, a neuroacanthocytosis syndrome with similar phenotypes to VPS13A-associated chorea-acanthocytosis (Roulis et al., 2018). The later age of onset for McLeod syndrome may be due to functional redundancy as the VPS13A binding site is shared with XK and one of its 8 homologs, XKR2 (Guillén-Samander et al., 2022). Identification of proteins in proximity to Vps13 at MCSs should uncover additional regulators that assist in lipid transfer. This could be performed with protein-proximity screens such as BioID, where proteins in the vicinity of the bait are biotinylated and subsequently 143 identified through mass spectrometry (Samavarchi-Tehrani et al., 2020), or through a protein complementation assay (PCA) based on the dihydrofolate reductase (DHFR) enzyme that provides drug resistance to methotrexate (Michnick et al., 2010). Multiple screens can be performed with Vps13 baits in cells overexpressing each of the known adaptors to determine which proteins are consistently found with Vps13 at all contact sites indicating a shared functional role, in comparison to those that are specific. One caveat is that tags on either end of Vps13 disrupt endogenous localization (Lang et al., 2015) and so the biotin ligase tag for BioID or split-DHFR tag for the PCA would need to be integrated in unconserved flexible loops that can be identified through inspection of AlphaFold models and multiple sequence alignments. Due to the large size of Vps13, two baits with tags integrated near N- or C-termini may be required to identify proteins in proximity at either end of the contact site. 5.5 The Repeating Beta Groove superfamily Along with others, we have identified Fmp27 and Hob2 as members of the newly classified repeating beta groove (RBG) family of bridging lipid transporters. These proteins form elongated lipid transfer channels comprised of multiple RBG modules containing 5 antiparallel ß-sheets followed by a disordered linker that frequently forms short ⍺-helices that lie overtop (Levine, 2022; Neuman et al., 2022b). The members in this MCS-localizing family include yeast Vps13 (VPS13A-D in humans), Atg2 (ATG2A-B in humans), Csf1 (BLTP1/KIAA1109 in humans) and paralogous Fmp27 and Hob2 (BLTP2/KIAA0100 in humans) (Fig 5.1), as well as mammalian BLTP3B/SHIP164 and its paralog BLTP3A/UHRF1BP1 (Neuman et al., 2022b; Levine, 2022). Of interest, the predicted channel length of each family member is variable with BLTP3B spanning ~12 nm while BLTP1 spans ~33 nm (Hanna et al., 2022; Levine, 2022; 144 Figure 5.1: RBG lipid transporters in budding yeast. Schematic of the five S. cerevisiae RBG proteins and their reported membrane contact site localizations. Insets show the known membrane targeting mechanisms used by each of the homologs. Vps13 is recruited to different contact sites by interactions with adaptor proteins mediated by its VAB domain. A predicted phospholipid-binding Pleckstrin homology (PH) domain may also contribute to membrane targeting in addition to membrane inserting amphipathic helices in the ATG2_C domain (Dall’Armellina et al., 2023), which is also found in Atg2. Atg2 is targeted to isolation membranes/phagophores through interactions with Atg18 and Atg9 (Gómez-Sánchez et al., 2018), and targets the ER with a conserved N-terminal helix in its chorein domain (Kotani et al., 2018b). The Fmp27, Hob2 and Csf1 proteins have N-terminal transmembrane ER anchors (Neuman et al., 2022a; Toulmay et al., 2022). Fmp27 and Hob2 are recruited to the PM through interactions with Hoi1. The putative direction of lipid transport is shown with dotted arrows within the lipid transfer channels. Only Csf1 has been proposed to transfer lipids towards the ER (Toulmay et al., 2022). Neuman et al., 2022b). The variation in length may be important for targeting specific MCS niches that may cluster related effectors together and exclude other MCS proteins. For example, VPS13A has been found to be recruited by XK to ER-PM contacts that are distinct from those 145 occupied by the ER-PM tether JPH3 (Guillén-Samander et al., 2022), and our data in yeast suggests Fmp27 occupies a distinct ER-PM contact site from the Tcb3 lipid transporter. In addition to variations in length, the different RBG family members are also suggested to form hydrophobic channels of varying widths. Vps13, Atg2 and BLTP3B have been demonstrated to transport and bind to numerous glycerophospholipids and do not appear to have specificity for lipid head groups (Kumar et al., 2018; Osawa et al., 2019; Hanna et al., 2022; Maeda et al., 2019; Li et al., 2020). Variations in channel width may provide differing specificities for lipid acyl chain lengths and/or saturation status between the different family members highlighting distinct functional roles. In fact, analysis of the lipidome of yeast cells lacking the ER-PM localized Csf1 identified an overall shortening of glycerophospholipid acyl chains with increased levels of unsaturation (John Peter et al., 2022). Models of Csf1 suggest it has one of the widest channels in yeast (Castro et al., 2022) and thus could transport bulkier unsaturated lipids between ER-PM contacts where they can be further modified. Of the RBG family members, only Vps13, Atg2 and BLTP3B have been partially or fully structurally resolved by cryo-EM or X-ray crystallography (Li et al., 2020; Kumar et al., 2018; Osawa et al., 2019; Valverde et al., 2019; Hanna et al., 2022). The remainder of the family members have instead been modeled with AlphaFold and trRosetta and thus cautious interpretation is needed. The large size of these proteins (>1,500 amino acids in yeast) makes computation of full-length models challenging and the overall architecture of the proteins can be skewed by regions modeled with low confidence. Such low confidence regions could include the unstructured loops between RBG domain ß-sheets, which are generally missing from experimentally determined crystal structures and are expected to be quite flexible (Osawa et al., 2019; Kumar et al., 2018). Negative-stain EM of Vps13 suggests the full-length structure is 146 relatively flexible (De et al., 2017) and such mobility cannot be captured by static AlphaFold models. Additionally, as the RBG family members are thought to bridge organelle membranes through membrane targeting determinants on either ends of their channels, structural predictions of these regions in the absence of membranes may not position them in orientations observed in vivo. Future cryo-EM and crystallographic reconstructions of the remainder of the RBG family members will be crucial for determining the channel lengths and widths of each member, and for developing improved in silico models to assess protein mobility and membrane interactions. A main outstanding question is what lipids are transferred by the RBG family. Promiscuous binding to glycerophospholipids has been determined for Vps13/VPS13D, Atg2 and BLTP3B (Valverde et al., 2019; Kumar et al., 2018; Hanna et al., 2022; Wang et al., 2021), however gel shift assays with purified Vps13 and Atg2 have also found weak binding to ceramide, the sphingolipid precursor, and cholesterol, and those with VPS13D also found binding to fatty acid moieties (Wang et al., 2021; Kumar et al., 2018; Valverde et al., 2019). To determine what lipids are transported by Fmp27 and Hob2, related in vitro gel shift assays with fluorescently labelled lipid species could be performed with purified N-terminal fragments. Additionally, affinity purification of these N-terminal fragments from cell lysates followed by liquid chromatography-tandem mass spectrometry (LC/MS/MS) may provide a better understanding of the lipids bound in vivo as similar experiments with the N-terminus of Vps13 suggest only glycerophospholipids truly bind (Kumar et al., 2018). Beyond lipid binding, it is important to determine if the associated lipids can be transported by Fmp27 and Hob2. Once candidate lipids are identified, in vitro FRET-based lipid transfer assays between liposomes should determine if these RBG family members are sufficient for their transport (Maeda et al., 2020; Kumar et al., 2018; Osawa et al., 2019; Valverde et al., 147 2019). To determine if lipid transport by Fmp27 and Hob2 is required for their function in ergosterol homeostasis, the distribution of mCherry-D4H can be evaluated in mutants with substitutions in a ring of hydrophobic residues within their lipid transfer channels, similar to the mutations tested in Vps13 (Li et al., 2020). The point mutations would need to be introduced into a background where the opposite paralog has been knocked out because deletion of both Fmp27 and Hob2 are needed to alter the distribution of accessible sterols. As mutation of the hydrophobic channels can result in misfolding, techniques such as DSLS that assess protein stability and folding (Senisterra et al., 2010; Kopec and Schneider, 2011) would be needed to ensure any null phenotypes are the direct result of a loss of lipid transfer. 5.6 Role of Hob proteins in lipid homeostasis and secretion We have determined in yeast that N-terminally ER-anchored Fmp27 is recruited to PM contact sites through interactions with Hoi1 for a shared function in the maintenance of ergosterol homeostasis. Curiously, unlike adaptors for Vps13 and Atg2, Hoi1 membrane localization is dependent on Fmp27. Using fluorescence microscopy-based chimeric recruitment assays guided by AlphaFold2 Multimer modeling of the Fmp27-Hoi1 interaction, we identified a large binding interface between a long ⍺-helical protrusion in Fmp27 and the C-terminus of Hoi1. Our work further suggests that in addition to this interface, a conserved helix in Hoi1 (residues 304-315) and a flexible loop in Fmp27 (residues 1791-1811) may contribute to binding but were unable to confidently model interactions at these sites. These regions could bind an additional protein that may bridge an interaction between Fmp27 and Hoi1 and may further help target the Fmp27-Hoi1 complex to the PM. 148 Like other RBG family proteins, the Fmp27 and Hob2 lipid channels are decorated with loops that may mediate protein-protein interactions with other effectors such as scramblases and biosynthetic enzymes. A PCA screen using split DHFR could be used to identify additional interactors and functionally related proteins in proximity to Fmp27-Hoi1 at ER-PM contact sites. Overexpressing Fmp27 baits with C-terminal split DHFR tags in cells overexpressing HOI1, where Fmp27 localizes to ER-PM contacts, or in HOI1 knockouts, where Fmp27 is diffuse in the ER, will allow proximal proteins specifically at PM contacts to be distinguished. Although we did not find a difference in localization when Fmp27 was C-terminally or internally tagged, we have not formally tested if these tags disrupt function, and it is possible that a C-terminal split DHFR tag may block some relevant protein interactions. To identify interactors in the ER, a related screen using an Fmp27 bait with an internal tag near the N-terminus would be required. Putative interactors identified in these screens can narrowed down to a candidate list by modeling them in complex with Fmp27 or Hoi1 using AlphaFold2 Multimer and validating confident interactions by co-immunoprecipitation. It is interesting to speculate that Laf1 and its paralog Dgr2 may physically interact with either Fmp27 or Hoi1. We found Laf1 is associated with both proteins in yeast chemicogenomics datasets (Hoepfner et al., 2014) and knockouts of HOI1 and LAF1 homologs in conjunction with a fibrillin mutation in C. elegans both result in an ingressed pharynx phenotype that is proposed to arise from secretion defects (Yochem et al., 2004; Kelley et al., 2015). Although yeast knockouts of HOI1 and LAF1 result in different sensitivities to the sterol-binding drug AmB and different mCherry-D4H distributions, this may be because interactions occur only under certain conditions. For example, in response to PM perturbing stresses TORC2-activated kinase Ypk1 phosphorylates Lam2/Lam4 resulting in the dissociation of Laf1 and Dgr2, thus inhibiting 149 retrograde sterol transport (Roelants et al., 2018; Topolska et al., 2020). It may be that Laf1/Dgr2, which remain localized at the PM (Topolska et al., 2020), can then interact with Fmp27 and Hoi1 and thus may have a broad function regulating ER-PM transport of sterols and sterol-shielding phospholipids. Co-immunoprecipitation experiments between Laf1 and Fmp27 or Hoi1 may identify an interaction, but likely only after activation of TORC2 signaling. The Fmp27 ⍺-helical extension and interacting Hoi1 helices are conserved in higher eukaryotes, indicating their homologs may similarly interact. Shared secretion defects observed in knockouts of the HOI1 homolog in fibrillin mutant nematodes, and in knockdowns of the FMP27 homolog in flies, further suggest these proteins may be functionally linked in higher eukaryotes. Determining if knockouts of the FMP27 homolog (F31C3.3) in C. elegans fibrillin mutants reproduce the pharynx malformations observed in related HOI1 homolog (sym-3) mutants will provide further evidence for a shared pathway for these homologs, as well as Laf1. Similar replication experiments can be performed in flies to determine if the small body size and regulated secretion defects observed in hobbit mutants are phenocopied by knockdowns of CG8671, the HOI1 homolog. Humans have a single Fmp27/Hob2 homologous protein known as BLTP2/KIAA0100 and two Hoi1 homologs, EEIG1/FAM102A and EEIG2/FAM102B. BLTP2 overexpression is implicated in cancers and BLTP2 has been demonstrated to function in cell aggregation, metastasis, invasion, and cell-cell adhesion in breast cancer lines (Song et al., 2006; Liu et al., 2014; Zhong et al., 2018). Exogenous expression of BLTP2 in fly hobbit mutants rescues body size and lethality suggesting it has a conserved function and likely localizes to ER-PM contacts (Neuman and Bashirullah, 2018). The Hoi1 homologs, EEIG1 and EEIG2, have been found to interact with a PM-localized protein or been shown to localize to the PM respectively (Popa et 150 al., 2020; Choi et al., 2013), supporting a possible function as the BLTP2 adaptor at this membrane. EEIG2 has recently been identified as necessary for the secretion of glycoproteins in a genome-wide CRISPR screen in HeLa cells (Popa et al., 2020). EEIG1 has instead been identified as an interactor for receptor activator of NF-κB (RANK) and is required for downstream signaling in osteoclast formation (Choi et al., 2013). Interestingly, both proteins are differentially expressed (Choi et al., 2013; Jeong et al., 2020; Wang et al., 2004) and thus may recruit the ER-localized BLTP2 to PM contact sites in response to different signals, acting as a switch to activate lipid transport at these sites. To determine if BLTP2 indeed interacts with EEIG1 and EEIG2, co-immunoprecipitation experiments can be performed in human cell lines. It will be particularly interesting to determine if putative interactions rely on the conserved ⍺-helical extension in BLTP2. If interactions are observed, fluorescent imaging experiments will be required to determine if these putative PM adaptors can recruit overexpressed BLTP2 to ER-PM contact sites. If a binding defective BLTP2 mutant can be identified, this mutant can be used to determine if lipid transport by BLTP2 is required for the RANK signaling and secretion functions of EEIG1 and EEIG2 respectively. For example, a similar secretion assay used to validate the role of EEIG2 in protein secretion can be used. In this assay, a construct consisting of a secretion signal fused to horse radish peroxidase (HRP) is expressed in HeLa cells and the extent of secretion determined by chemiluminescence (Popa et al., 2020). If a BLTP2 mutant that is unable to bind EEIG2 reconstitutes the EEIG2 knockdown secretion defect this could indicate that lipid transfer is required for this function. Furthermore, this would confirm a shared functional role in secretion for mammalian Fmp27 and Hoi1 homologs. 151 5.7 Limitations and Future Directions The work described here contributes to our knowledge of the membrane targeting strategies of the RBG protein superfamily and identifies a PM lipid homeostasis function for Fmp27 and Hob2, newly identified yeast family members. The interpretation of these findings and their extension to other RBG family members is subject to certain caveats, and key outstanding questions in the field remain. We have taken advantage of the budding yeast model to identify conserved domains required for RBG protein-adaptor interactions and for RBG protein recruitment to membranes. Although the adaptor binding and membrane localization functions of human VPS13 VAB domains have been validated by others (Kumar et al., 2018; Wang et al., 2021; Guillén-Samander et al., 2021; Tornero-Écija et al., 2023), the relevance of the identified interaction sites in yeast Fmp27/Hob2 and their PM interactor, Hoi1, in the homologous human proteins remains to be tested. Further, our work identifying Hoi1 as the PM adaptor and determining the minimal Fmp27-Hoi1 interaction interface is based on co-immunoprecipitation and fluorescent microscopy recruitment assays which cannot confirm direct interactions. Validation of the predicted Fmp27-Hoi1 interface by structural reconstruction, such as the X-crystallography validation of the Vps13 VAB-PxP interaction (Adlakha et al., 2022), will be critical for confirming direct binding and validating the extensive interaction interface predicted by AlphaFold2 Multimer. The redistribution of ergosterol seen in FMP27 HOB2 or HOI1 mutants is proposed to result from a loss of ER-PM lipid transport, however direct evidence for a lipid transport function for paralogous Fmp27 and Hob2 is missing. Biochemical assays to determine the lipid binding specificity and lipid transfer function of these proteins, such as fluorescent gel shift assays and FRET-based liposome transfer assays, are required. Once lipid transport is confirmed 152 in vitro, the necessity for this function in the ergosterol distribution phenotype can be assessed with a lipid transfer defective Fmp27 mutant that blocks the transport channel, similar to tested mutations in Vps13 (Li et al., 2020), in a hob2Δ mutant. Design of this lipid transfer defective mutant would be improved following structural characterization of the full-length Fmp27 or Hob2 proteins. Currently only Vps13, Atg2 and BLTP3B have been partially or fully structurally resolved (Li et al., 2020; Kumar et al., 2018; Valverde et al., 2019; Hanna et al., 2022; Osawa et al., 2019) and thus the classification of the majority of the RBG superfamily members is based on structural predictions and predicted remote structural homology searches. As the field progresses, structural characterization of these family members will be crucial for determining the differences in channel lengths and widths between family members which can indicate different contact site or lipid specificities. Cryo-EM reconstructions of the full-length proteins bound to liposomes can further contribute to our knowledge of the flexibility of these proteins and their dynamic association with membranes. Future work identifying proteins in proximity to the RBG family members will likely add to our understanding of the drivers of lipid transport as confirmation of associations with lipid transporters, such as scramblases, or lipid biosynthetic enzymes can aid in the development of better in vitro and in silico models of lipid transport by members of this family. 5.8 Concluding Remarks The RBG superfamily of proteins form newly characterized lipid transfer channels that physically bridge organelles at MCSs. Mutations in the various human family members are associated with different neurological disorders as well as cancer (Seong et al., 2018; Rampoldi et al., 2001; Kolehmainen et al., 2003; Lesage et al., 2016; Jansen et al., 2017; Gueneau et al., 153 2018), highlighting the important functions of this family. Understanding how these proteins target membranes is essential for determining which contact sites are occupied by each family member, and how mutations in these proteins result in distinct diseases. In this dissertation, we identify a previously uncharacterized domain in VPS13 proteins which binds to organelle-specific adaptors and is required for membrane recruitment in budding yeast. Our studies proposed a single adaptor binding site in this domain which is blocked by a disease-associated substitution, suggesting a conserved adaptor binding function for this domain in human VPS13 proteins which has now been confirmed by others (Kumar et al., 2018; Guillén-Samander et al., 2021; Tornero-Écija et al., 2023). We additionally independently identified Fmp27 and Hob2 as novel RBG superfamily members with a role in ergosterol homeostasis, and show they similarly rely on adaptor interactions for recruitment to contact sites. Future work will be required to determine what lipids are transferred by the various RBG family members and where they localize in humans, particularly in response to cellular cues or in different cell types. Understanding what contact sites are occupied by each member and what lipids are transported will be critical for developing a better understanding of the associated disease pathologies. 154 Bibliography Adelmant, G., B. Garg, M. Tavares, J.D. Card, and J.A. Marto. 2019. Tandem Affinity Purification and Mass Spectrometry (TAP-MS) for the Analysis of Protein Complexes. Curr Protoc Protein Sci. 96:e84. doi:10.1002/cpps.84. Adlakha, J., Z. Hong, P. Li, and K.M. Reinisch. 2022. Structural and biochemical insights into lipid transport by VPS13 proteins. Journal of Cell Biology. 221:e202202030. doi:10.1083/jcb.202202030. Alberts, B., A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter. 2002a. The Compartmentalization of Cells. Molecular Biology of the Cell. 4th edition. Alberts, B., A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter. 2002b. The Lipid Bilayer. Molecular Biology of the Cell. 4th edition. Alberts, B., A. Johnson, J. Lewis, M. Raff, K. Roberts, and P. Walter. 2002c. Intracellular Vesicular Traffic. Molecular Biology of the Cell. 4th edition. Alpy, F., and C. Tomasetto. 2014. START ships lipids across interorganelle space. Biochimie. 96:85–95. doi:10.1016/j.biochi.2013.09.015. Ammar, M.R., N. Kassas, S. Chasserot-Golaz, M.-F. Bader, and N. Vitale. 2013. Lipids in Regulated Exocytosis: What are They Doing? Front. Endocrinol. 4. doi:10.3389/fendo.2013.00125. Anding, A.L., C. Wang, T.-K. Chang, D.A. Sliter, C.M. Powers, K. Hofmann, R.J. Youle, and E.H. Baehrecke. 2018. Vps13D encodes a ubiquitin binding protein that is required for the regulation of mitochondrial size and clearance. Current Biology. 28:287-295.e6. doi:10.1016/j.cub.2017.11.064. Andrejeva, G., S. Gowan, G. Lin, A.C.L. Wong Te Fong, E. Shamsaei, H.G. Parkes, J. Mui, F.I. Raynaud, Y. Asad, G. Vizcay-Barrena, J. Nikitorowicz-Buniak, M. Valenti, L. Howell, R.A. Fleck, L.A. Martin, V. Kirkin, M.O. Leach, and Y.L. Chung. 2019. De novo phosphatidylcholine synthesis is required for autophagosome membrane formation and maintenance during autophagy. Autophagy. 1–17. doi:10.1080/15548627.2019.1659608. Arora, A., J.H. Taskinen, and V.M. Olkkonen. 2022. Coordination of inter-organelle communication and lipid fluxes by OSBP-related proteins. Prog Lipid Res. 86:101146. doi:10.1016/j.plipres.2022.101146. Ashburner, M., C.A. Ball, J.A. Blake, D. Botstein, H. Butler, J.M. Cherry, A.P. Davis, K. Dolinski, S.S. Dwight, J.T. Eppig, M.A. Harris, D.P. Hill, L. Issel-Tarver, A. Kasarskis, S. Lewis, J.C. Matese, J.E. Richardson, M. Ringwald, G.M. Rubin, and G. Sherlock. 2000. Gene Ontology: tool for the unification of biology. Nat Genet. 25:25–29. doi:10.1038/75556. 155 Ashkenazy, H., S. Abadi, E. Martz, O. Chay, I. Mayrose, T. Pupko, and N. Ben-Tal. 2016. ConSurf 2016: an improved methodology to estimate and visualize evolutionary conservation in macromolecules. Nucleic Acids Res. 44:W344–W350. doi:10.1093/nar/gkw408. Bailey, T.L., and C. Elkan. 1994. Fitting a mixture model by expectation maximization to discover motifs in biopolymers. Proceedings of the Second International Conference on Intelligent Systems for Molecular Biology. 28–36. Baldwin, H.A., C. Wang, G. Kanfer, H.V. Shah, A. Velayos-Baeza, M. Dulovic-Mahlow, N. Brüggemann, A. Anding, E.H. Baehrecke, D. Maric, W.A. Prinz, and R.J. Youle. 2021. VPS13D promotes peroxisome biogenesis. Journal of Cell Biology. 220:e202001188. doi:10.1083/jcb.202001188. Balicza, P., N.Á. Varga, B. Bolgár, K. Pentelényi, R. Bencsik, A. Gál, A. Gézsi, C. Prekop, V. Molnár, and M.J. Molnár. 2019. Comprehensive Analysis of Rare Variants of 101 Autism-Linked Genes in a Hungarian Cohort of Autism Spectrum Disorder Patients. Frontiers in Genetics. 10:434. doi:10.3389/fgene.2019.00434. Balikova, I., AE. Lehesjoki, T.J. de Ravel, B. Thienpont, K.E. Chandler, J. Clayton-Smith, AL. Traskelin, JP. Fryns, and J.R. Vermeesch. 2009. Deletions in the VPS13B (COH1) gene as a cause of Cohen syndrome. Human Mutation. 30:E845–E854. doi:10.1002/humu.21065. Balla, T., N. Sengupta, and Y.J. Kim. 2020. Lipid synthesis and transport are coupled to regulate membrane lipid dynamics in the endoplasmic reticulum. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1865:158461. doi:10.1016/j.bbalip.2019.05.005. Ballweg, S., and R. Ernst. 2017. Control of membrane fluidity: the OLE pathway in focus. Biological Chemistry. 398:215–228. doi:10.1515/hsz-2016-0277. Bankaitis, V.A., L.M. Johnson, and S.D. Emr. 1986. Isolation of yeast mutants defective in protein targeting to the vacuole. Proceedings of the National Academy of Sciences. 83:9075–9. doi:10.1073/PNAS.83.23.9075. Baskaran, S., M.J. Ragusa, E. Boura, and J.H. Hurley. 2012. Two-site recognition of phosphatidylinositol 3-phosphate by PROPPINs in autophagy. Molecular Cell. 47:339–348. doi:10.1016/j.molcel.2012.05.027. Baumann, N.A., D.P. Sullivan, H. Ohvo-Rekilä, C. Simonot, A. Pottekat, Z. Klaassen, C.T. Beh, and A.K. Menon. 2005. Transport of Newly Synthesized Sterol to the Sterol-Enriched Plasma Membrane Occurs via Nonvesicular Equilibration. Biochemistry. 44:5816–5826. doi:10.1021/bi048296z. 156 Bean, B.D.M., S.K. Dziurdzik, K.L. Kolehmainen, C.M.S. Fowler, W.K. Kwong, L.I. Grad, M. Davey, C. Schluter, and E. Conibear. 2018. Competitive organelle-specific adaptors recruit Vps13 to membrane contact sites. Journal of Cell Biology. 217:3593–3607. doi:10.1083/jcb.201804111. Bean, B.D.M., M. Whiteway, and V.J.J. Martin. 2022. The MyLO CRISPR-Cas9 toolkit: a markerless yeast localization and overexpression CRISPR-Cas9 toolkit. G3 Genes|Genomes|Genetics. 12:jkac154. doi:10.1093/g3journal/jkac154. Behnia, R., and S. Munro. 2005. Organelle identity and the signposts for membrane traffic. Nature. 438:597–604. doi:10.1038/nature04397. Besprozvannaya, M., E. Dickson, H. Li, K.S. Ginburg, D.M. Bers, J. Auwerx, and J. Nunnari. 2018. GRAM domain proteins specialize functionally distinct ER-PM contact sites in human cells. eLife. 7:e31019. doi:10.7554/eLife.31019. Bevers, E.M., and P.L. Williamson. 2010. Phospholipid scramblase: An update. FEBS Letters. 584:2724–2730. doi:10.1016/j.febslet.2010.03.020. Bian, X., Y. Saheki, and P. De Camilli. 2018. Ca2+ releases E‐Syt1 autoinhibition to couple ER‐plasma membrane tethering with lipid transport. The EMBO Journal. 37:219–234. doi:10.15252/embj.201797359. Bieberich, E. 2018. Sphingolipids and lipid rafts: Novel concepts and methods of analysis. Chem Phys Lipids. 216:114–131. doi:10.1016/j.chemphyslip.2018.08.003. Biegert, A., and J. Söding. 2008. De novo identification of highly diverged protein repeats by probabilistic consistency. Bioinformatics. 24:807–814. doi:10.1093/bioinformatics/btn039. Blom, T., P. Somerharju, and E. Ikonen. 2011. Synthesis and Biosynthetic Trafficking of Membrane Lipids. Cold Spring Harb Perspect Biol. 3:a004713. doi:10.1101/cshperspect.a004713. Blomberg, N., E. Baraldi, M. Nilges, and M. Saraste. 1999. The PH superfold: A structural scaffold for multiple functions. Trends in Biochemical Sciences. 24:441–445. doi:10.1016/S0968-0004(99)01472-3. Bogdanov, M., W. Dowhan, and H. Vitrac. 2014. Lipids and topological rules governing membrane protein assembly. Biochim Biophys Acta. 1843:1475–1488. doi:10.1016/j.bbamcr.2013.12.007. Bonifacino, J.S., and B.S. Glick. 2004. The Mechanisms of Vesicle Budding and Fusion. Cell. 116:153–166. doi:10.1016/S0092-8674(03)01079-1. 157 Bowsher, C.G., and A.K. Tobin. 2001. Compartmentation of metabolism within mitochondria and plastids. Journal of Experimental Botany. 52:513–527. doi:10.1093/jexbot/52.356.513. Brickner, J.H., and R.S. Fuller. 1997. SOI1 Encodes a Novel, Conserved Protein That Promotes TGN–Endosomal Cycling of Kex2p and Other Membrane Proteins by Modulating the Function of Two TGN Localization Signals. Journal of Cell Biology. 139:23–36. Brückner, A., C. Polge, N. Lentze, D. Auerbach, and U. Schlattner. 2009. Yeast Two-Hybrid, a Powerful Tool for Systems Biology. Int J Mol Sci. 10:2763–2788. doi:10.3390/ijms10062763. Calianese, D.C., and R.B. Birge. 2020. Biology of phosphatidylserine (PS): basic physiology and implications in immunology, infectious disease, and cancer. Cell Communication and Signaling. 18:41. doi:10.1186/s12964-020-00543-8. Calzada, E., E. Avery, P.N. Sam, A. Modak, C. Wang, J.M. McCaffery, X. Han, N.N. Alder, and S.M. Claypool. 2019. Phosphatidylethanolamine made in the inner mitochondrial membrane is essential for yeast cytochrome bc1 complex function. Nat Commun. 10:1432. doi:10.1038/s41467-019-09425-1. Castro, I.G., S.P. Shortill, S.K. Dziurdzik, A. Cadou, S. Ganesan, R. Valenti, Y. David, M. Davey, C. Mattes, F.B. Thomas, R.E. Avraham, H. Meyer, A. Fadel, E.J. Fenech, R. Ernst, V. Zaremberg, T.P. Levine, C. Stefan, E. Conibear, and M. Schuldiner. 2022. Systematic analysis of membrane contact sites in Saccharomyces cerevisiae uncovers modulators of cellular lipid distribution. eLife. 11:e74602. doi:10.7554/eLife.74602. Chamberlain, L.H., R.D. Burgoyne, and G.W. Gould. 2001. SNARE proteins are highly enriched in lipid rafts in PC12 cells: Implications for the spatial control of exocytosis. Proceedings of the National Academy of Sciences. 98:5619–5624. doi:10.1073/pnas.091502398. Chasserot-Golaz, S., J.R. Coorssen, F.A. Meunier, and N. Vitale. 2010. Lipid Dynamics in Exocytosis. Cell Mol Neurobiol. 30:1335–1342. doi:10.1007/s10571-010-9577-x. Chen, S., M. Mari, S. Parashar, D. Liu, Y. Cui, F. Reggiori, P.J. Novick, and S. Ferro-Novick. 2020. Vps13 is required for the packaging of the ER into autophagosomes during ER-phagy. Proceedings of the National Academy of Sciences. 117:18530–18539. doi:10.1073/pnas.2008923117. Choi, H.K., H.R. Kang, E. Jung, T.E. Kim, J.J. Lin, and S.Y. Lee. 2013. Early estrogen-induced gene 1, a novel RANK signaling component, is essential for osteoclastogenesis. Cell Research. 23:524–536. doi:10.1038/cr.2013.33. Chowdhury, S., C. Otomo, A. Leitner, K. Ohashi, R. Aebersold, G.C. Lander, and T. Otomo. 2018. Insights into autophagosome biogenesis from structural and biochemical analyses 158 of the ATG2A-WIPI4 complex. Proceedings of the National Academy of Sciences. 115:9792–9801. doi:10.1073/PNAS.1811874115. Churchward, M.A., T. Rogasevskaia, J. Höfgen, J. Bau, and J.R. Coorssen. 2005. Cholesterol facilitates the native mechanism of Ca2+-triggered membrane fusion. Journal of Cell Science. 118:4833–4848. doi:10.1242/jcs.02601. Collado, J., M. Kalemanov, F. Campelo, C. Bourgoint, F. Thomas, R. Loewith, A. Martínez-Sánchez, W. Baumeister, C.J. Stefan, and R. Fernández-Busnadiego. 2019. Tricalbin-Mediated Contact Sites Control ER Curvature to Maintain Plasma Membrane Integrity. Developmental Cell. 51:476-487.e7. doi:10.1016/j.devcel.2019.10.018. Cowart, L.A., and L.M. Obeid. 2007. Yeast Sphingolipids: Recent developments in understanding biosynthesis, regulation, and function. Biochim Biophys Acta. 1771:421–431. doi:10.1016/j.bbalip.2006.08.005. Cui, L., H. Li, Y. Xi, Q. Hu, H. Liu, J. Fan, Y. Xiang, X. Zhang, W. Shui, and Y. Lai. 2022. Vesicle trafficking and vesicle fusion: mechanisms, biological functions, and their implications for potential disease therapy. Mol Biomed. 3:29. doi:10.1186/s43556-022-00090-3. Da Costa, R., M. Bordessoules, M. Guilleman, V. Carmignac, V. Lhussiez, H. Courot, A. Bataille, A. Chlémaire, C. Bruno, P. Fauque, C. Thauvin, L. Faivre, and L. Duplomb. 2020. Vps13b is required for acrosome biogenesis through functions in Golgi dynamic and membrane trafficking. Cellular and Molecular Life Sciences. 77:511–529. doi:10.1007/s00018-019-03192-4. Dall’Armellina, F., M. Stagi, and L.E. Swan. 2023. In silico modeling human VPS13 proteins associated with donor and target membranes suggests lipid transfer mechanisms. Proteins: Structure, Function, and Bioinformatics. 91:439–455. doi:10.1002/prot.26446. Dalton, L.E., B.D.M. Bean, M. Davey, and E. Conibear. 2017. Quantitative high-content imaging identifies novel regulators of Neo1 trafficking at endosomes. MBoC. 28:1539–1550. doi:10.1091/mbc.e16-11-0772. Datta, S., Y. Liu, H. Hariri, J. Bowerman, and W.M. Henne. 2019. Cerebellar ataxia disease-associated Snx14 promotes lipid droplet growth at ER-droplet contacts. Journal of Cell Biology. 218:1335–1351. doi:10.1083/jcb.201808133. Day, K.J., and J.C. Stachowiak. 2020. Biophysical Forces in Membrane Bending and Traffic. Curr Opin Cell Biol. 65:72–77. doi:10.1016/j.ceb.2020.02.017. De, M., A.N. Oleskie, M. Ayyash, S. Dutta, L. Mancour, M.E. Abazeed, E.J. Brace, G. Skiniotis, and R.S. Fuller. 2017. The Vps13p-Cdc31p complex is directly required for TGN late endosome transport and TGN homotypic fusion. Journal of Cell Biology. 216:425–439. doi:10.1083/jcb.201606078. 159 Di Mattia, T., A. Martinet, S. Ikhlef, A.G. McEwen, Y. Nominé, C. Wendling, P. Poussin‐Courmontagne, L. Voilquin, P. Eberling, F. Ruffenach, J. Cavarelli, J. Slee, T.P. Levine, G. Drin, C. Tomasetto, and F. Alpy. 2020. FFAT motif phosphorylation controls formation and lipid transfer function of inter‐organelle contacts. The EMBO Journal. 39:e104369. doi:10.15252/embj.2019104369. Dickson, E.J., and B. Hille. 2019. Understanding phosphoinositides: rare, dynamic, and essential membrane phospholipids. Biochem J. 476:1–23. doi:10.1042/BCJ20180022. Dobson-Stone, C., A. Danek, L. Rampoldi, R. Hardie, R. Chalmers, N. Wood, S. Bohlega, M. Dotti, A. Federico, M. Shizuka, M. Tanaka, M. Watanabe, Y. Ikeda, M. Brin, L. Goldfarb, B. Karp, S. Mohiddin, L. Fananapazir, A. Storch, A. Fryer, P. Maddison, I. Sibon, P. Trevisol-Bittencourt, C. Singer, I. Caballero, J. Aasly, K. Schmierer, R. Dengler, L.-P. Hiersemenzel, M. Zeviani, V. Meiner, A. Lossos, S. Johnson, F. Mercado, G. Sorrentino, N. Dupré, G. Rouleau, J. Volkmann, J. Arpa, A. Lees, G. Geraud, S. Chouinard, A. Németh, and A. Monaco. 2002. Mutational spectrum of the CHAC gene in patients with chorea-acanthocytosis. European Journal of Human Genetics. 10:773–781. doi:10.1038/sj.ejhg.5200866. Drin, G., J.-F. Casella, R. Gautier, T. Boehmer, T.U. Schwartz, and B. Antonny. 2007. A general amphipathic α-helical motif for sensing membrane curvature. Nature Structural & Molecular Biology. 14:138–146. doi:10.1038/nsmb1194. Drin, G., J.M. von Filseck, and A. Čopič. 2016. New molecular mechanisms of inter-organelle lipid transport. Biochemical Society Transactions. 44:486–492. doi:10.1042/BST20150265. Du, Z., H. Su, W. Wang, L. Ye, H. Wei, Z. Peng, I. Anishchenko, D. Baker, and J. Yang. 2021. The trRosetta server for fast and accurate protein structure prediction. Nat Protoc. 16:5634–5651. doi:10.1038/s41596-021-00628-9. Dupont, N., S. Chauhan, J. Arko-Mensah, E.F. Castillo, A. Masedunskas, R. Weigert, H. Robenek, T. Proikas-Cezanne, and V. Deretic. 2014. Neutral lipid stores and lipase PNPLA5 contribute to autophagosome biogenesis. Current Biology. 24:609–620. doi:10.1016/j.cub.2014.02.008. Dziurdzik, S.K., B.D.M. Bean, M. Davey, and E. Conibear. 2020. A VPS13D spastic ataxia mutation disrupts the conserved adaptor-binding site in yeast Vps13. Human Molecular Genetics. 29:635–648. doi:10.1093/hmg/ddz318. Dziurdzik, S.K., and E. Conibear. 2021. The Vps13 Family of Lipid Transporters and Its Role at Membrane Contact Sites. International Journal of Molecular Sciences 2021, Vol. 22, Page 2905. 22:2905. doi:10.3390/IJMS22062905. Edgar, R.C. 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research. 32:1792–1797. doi:10.1093/nar/gkh340. 160 Egea, P.F. 2021. Mechanisms of Non-Vesicular Exchange of Lipids at Membrane Contact Sites: Of Shuttles, Tunnels and, Funnels. Frontiers in Cell and Developmental Biology. 9. Eisenberg-Bord, M., N. Shai, M. Schuldiner, and M. Bohnert. 2016. A Tether Is a Tether Is a Tether: Tethering at Membrane Contact Sites. Developmental Cell. 39:395–409. doi:10.1016/j.devcel.2016.10.022. Elbaz-Alon, Y., M. Eisenberg-Bord, V. Shinder, S.B. Stiller, E. Shimoni, N. Wiedemann, T. Geiger, and M. Schuldiner. 2015. Lam6 Regulates the Extent of Contacts between Organelles. Cell Reports. 12:7–14. doi:10.1016/j.celrep.2015.06.022. Elbaz-Alon, Y., E. Rosenfeld-Gur, V. Shinder, A.H. Futerman, T. Geiger, and M. Schuldiner. 2014. A dynamic interface between vacuoles and mitochondria in yeast. Dev Cell. 30:95–102. doi:10.1016/j.devcel.2014.06.007. Fidler, D.R., S.E. Murphy, K. Courtis, P. Antonoudiou, R. El-Tohamy, J. Ient, and T.P. Levine. 2016. Using HHsearch to tackle proteins of unknown function: A pilot study with PH domains. Traffic. 17:1214–1226. doi:10.1111/tra.12432. Föller, M., A. Hermann, S. Gu, I. Alesutan, S.M. Qadri, O. Borst, E. Schmidt, F. Schiele, J. Mu, C. Saft, L. Schöls, H. Lerche, C. Stournaras, A. Storch, and F. Lang. 2012. Chorein-sensitive polymerization of cortical actin and suicidal cell death in chorea-acanthocytosis. FASEB Journal. 26:1526–1534. doi:10.1096/fj.11-198317. Gao, K., R. Oerlemans, and M.R. Groves. 2020. Theory and applications of differential scanning fluorimetry in early-stage drug discovery. Biophys Rev. 12:85–104. doi:10.1007/s12551-020-00619-2. Gatta, A.T., and T.P. Levine. 2017. Piecing Together the Patchwork of Contact Sites. Trends in Cell Biology. 27:214–229. doi:10.1016/j.tcb.2016.08.010. Gatta, A.T., L.H. Wong, Y.Y. Sere, D.M. Calderón-Noreña, S. Cockcroft, A.K. Menon, and T.P. Levine. 2015. A new family of StART domain proteins at membrane contact sites has a role in ER-PM sterol transport. eLife. 4:e07253. doi:10.7554/eLife.07253. Gault, C., L. Obeid, and Y. Hannun. 2010. An overview of sphingolipid metabolism: from synthesis to breakdown. Adv Exp Med Biol. 688:1–23. Gauthier, J., I.A. Meijer, D. Lessel, N.E. Mencacci, D. Krainc, M. Hempel, K. Tsiakas, H. Prokisch, E. Rossignol, M.H. Helm, L.H. Rodan, J. Karamchandani, M. Carecchio, S.J. Lubbe, A. Telegrafi, L.B. Henderson, K. Lorenzo, S.E. Wallace, I.A. Glass, F.F. Hamdan, J.L. Michaud, G.A. Rouleau, and P.M. Campeau. 2018. Recessive mutations in VPS13D cause childhood onset movement disorders. Annals of Neurology. 83:1089–1095. doi:10.1002/ana.25204. Gerace, E., and D. Moazed. 2015. Affinity Purification of Protein Complexes Using TAP Tags. Methods Enzymol. 559:37–52. doi:10.1016/bs.mie.2014.11.007. 161 Ghanbarpour, A., D.P. Valverde, T.J. Melia, and K.M. Reinisch. 2021. A model for a partnership of lipid transfer proteins and scramblases in membrane expansion and organelle biogenesis. Proceedings of the National Academy of Sciences. 118:e2101562118. doi:10.1073/pnas.2101562118. Giaever, G., A.M. Chu, L. Ni, C. Connelly, L. Riles, S. Véronneau, S. Dow, A. Lucau-Danila, K. Anderson, B. André, A.P. Arkin, A. Astromoff, M. El-Bakkoury, R. Bangham, R. Benito, S. Brachat, S. Campanaro, M. Curtiss, K. Davis, A. Deutschbauer, K.-D. Entian, P. Flaherty, F. Foury, D.J. Garfinkel, M. Gerstein, D. Gotte, U. Güldener, J.H. Hegemann, S. Hempel, Z. Herman, D.F. Jaramillo, D.E. Kelly, S.L. Kelly, P. Kötter, D. LaBonte, D.C. Lamb, N. Lan, H. Liang, H. Liao, L. Liu, C. Luo, M. Lussier, R. Mao, P. Menard, S.L. Ooi, J.L. Revuelta, C.J. Roberts, M. Rose, P. Ross-Macdonald, B. Scherens, G. Schimmack, B. Shafer, D.D. Shoemaker, S. Sookhai-Mahadeo, R.K. Storms, J.N. Strathern, G. Valle, M. Voet, G. Volckaert, C. Wang, T.R. Ward, J. Wilhelmy, E.A. Winzeler, Y. Yang, G. Yen, E. Youngman, K. Yu, H. Bussey, J.D. Boeke, M. Snyder, P. Philippsen, R.W. Davis, and M. Johnston. 2002. Functional profiling of the Saccharomyces cerevisiae genome. Nature. 418:387–391. doi:10.1038/nature00935. Gillingham, A.K., R. Sinka, I.L. Torres, K.S. Lilley, and S. Munro. 2014. Toward a comprehensive map of the effectors of rab GTPases. Developmental Cell. 31:358–373. doi:10.1016/j.devcel.2014.10.007. Giménez-Andrés, M., A. Čopič, and B. Antonny. 2018. The many faces of amphipathic helices. Biomolecules. 8. doi:10.3390/biom8030045. Gómez-Sánchez, R., J. Rose, R. Guimarães, M. Mari, D. Papinski, E. Rieter, W.J. Geerts, R. Hardenberg, C. Kraft, C. Ungermann, and F. Reggiori. 2018. Atg9 establishes Atg2-dependent contact sites between the endoplasmic reticulum and phagophores. Journal of Cell Biology. 217:2743–2763. doi:10.1083/jcb.201710116. González Montoro, A., K. Auffarth, C. Hönscher, M. Bohnert, T. Becker, B. Warscheid, F. Reggiori, M. van der Laan, F. Fröhlich, and C. Ungermann. 2018. Vps39 Interacts with Tom40 to Establish One of Two Functionally Distinct Vacuole-Mitochondria Contact Sites. Developmental Cell. 45:621-636.e7. doi:10.1016/j.devcel.2018.05.011. Graham, T.R. 2004. Flippases and vesicle-mediated protein transport. Trends in Cell Biology. 14:670–677. doi:10.1016/j.tcb.2004.10.008. Grant, C.E., T.L. Bailey, and W.S. Noble. 2017. FIMO: scanning for occurrences of a given motif. Bioinformatics. 27:1017–1018. doi:10.1093/bioinformatics/btr064. Gray, K.C., D.S. Palacios, I. Dailey, M.M. Endo, B.E. Uno, B.C. Wilcock, and M.D. Burke. 2012. Amphotericin primarily kills yeast by simply binding ergosterol. Proceedings of the National Academy of Sciences. 109:2234–2239. doi:10.1073/pnas.1117280109. 162 Grimm, J., H. Shi, W. Wang, A.M. Mitchell, N.S. Wingreen, K.C. Huang, and T.J. Silhavy. 2020. The inner membrane protein YhdP modulates the rate of anterograde phospholipid flow in Escherichia coli. Proceedings of the National Academy of Sciences. 117:26907–26914. doi:10.1073/pnas.2015556117. Guan, X.L., C.M. Souza, H. Pichler, G. Dewhurst, O. Schaad, K. Kajiwara, H. Wakabayashi, T. Ivanova, G.A. Castillon, M. Piccolis, F. Abe, R. Loewith, K. Funato, M.R. Wenk, and H. Riezman. 2009. Functional Interactions between Sphingolipids and Sterols in Biological Membranes Regulating Cell Physiology. MBoC. 20:2083–2095. doi:10.1091/mbc.e08-11-1126. Gueneau, L., R.J. Fish, H.E. Shamseldin, N. Voisin, F. Tran Mau-Them, E. Preiksaitiene, G.R. Monroe, A. Lai, A. Putoux, F. Allias, Q. Ambusaidi, L. Ambrozaityte, L. Cimbalistienė, J. Delafontaine, N. Guex, M. Hashem, W. Kurdi, S.S. Jamuar, L.J. Ying, C. Bonnard, T. Pippucci, S. Pradervand, B. Roechert, P.M. van Hasselt, M. Wiederkehr, C.F. Wright, I. Xenarios, G. van Haaften, C. Shaw-Smith, E.M. Schindewolf, M. Neerman-Arbez, D. Sanlaville, G. Lesca, L. Guibaud, B. Reversade, J. Chelly, V. Kučinskas, F.S. Alkuraya, and A. Reymond. 2018. KIAA1109 Variants Are Associated with a Severe Disorder of Brain Development and Arthrogryposis. Am J Hum Genet. 102:116–132. doi:10.1016/j.ajhg.2017.12.002. Guillén-Samander, A., M. Leonzino, M.G. Hanna IV, N. Tang, H. Shen, and P. De Camilli. 2021. VPS13D bridges the ER to mitochondria and peroxisomes via Miro. Journal of Cell Biology. 220:e202010004. doi:10.1083/jcb.202010004. Guillén-Samander, A., Y. Wu, S.S. Pineda, F.J. García, J.N. Eisen, M. Leonzino, B. Ugur, M. Kellis, M. Heiman, and P. De Camilli. 2022. A partnership between the lipid scramblase XK and the lipid transfer protein VPS13A at the plasma membrane. Proceedings of the National Academy of Sciences. 119:e2205425119. doi:10.1073/pnas.2205425119. Hanna, M., A. Guillén-Samander, and P. De Camilli. 2023. RBG Motif Bridge-Like Lipid Transport Proteins: Structure, Functions, and Open Questions. Annu Rev Cell Dev Biol. doi:10.1146/annurev-cellbio-120420-014634. Hanna, M.G., P.H. Suen, Y. Wu, K.M. Reinisch, and P. De Camilli. 2022. SHIP164 is a chorein motif lipid transfer protein that controls endosome–Golgi membrane traffic. Journal of Cell Biology. 221:e202111018. doi:10.1083/jcb.202111018. Hariri, H., S. Rogers, R. Ugrankar, Y.L. Liu, J.R. Feathers, and W.M. Henne. 2018. Lipid droplet biogenesis is spatially coordinated at ER-vacuole contacts under nutritional stress. EMBO Reports. 19:57–72. doi:10.15252/embr.201744815. Hariri, H., N. Speer, J. Bowerman, S. Rogers, G. Fu, E. Reetz, S. Datta, J.R. Feathers, R. Ugrankar, D. Nicastro, and W.M. Henne. 2019. Mdm1 maintains endoplasmic reticulum homeostasis by spatially regulating lipid droplet biogenesis. Journal of Cell Biology. 218:1319–1334. doi:10.1083/jcb.201808119. 163 Heino, S., S. Lusa, P. Somerharju, C. Ehnholm, V.M. Olkkonen, and E. Ikonen. 2000. Dissecting the role of the Golgi complex and lipid rafts in biosynthetic transport of cholesterol to the cell surface. Proceedings of the National Academy of Sciences. 97:8375–8380. Helle, S.C.J., G. Kanfer, K. Kolar, A. Lang, A.H. Michel, and B. Kornmann. 2013. Organization and function of membrane contact sites. Biochim Biophys Acta. 1833:2526–2541. doi:10.1016/j.bbamcr.2013.01.028. Henne, W.M. 2017. Discovery and Roles of ER-Endolysosomal Contact Sites in Disease. Advances in Experimental Medicine and Biology. 997:135–147. doi:10.1007/978-981-10-4567-7_10. Henne, W.M., L. Zhu, Z. Balogi, C. Stefan, J.A. Pleiss, and S.D. Emr. 2015. Mdm1/Snx13 is a novel ER-endolysosomal interorganelle tethering protein. Journal of Cell Biology. 210:541–551. doi:10.1083/jcb.201503088. Herdman, C., and T. Moss. 2016. Extended-Synaptotagmins (E-Syts); the extended story. Pharmacological Research. 107:48–56. doi:10.1016/j.phrs.2016.01.034. Hoepfner, D., S.B. Helliwell, H. Sadlish, S. Schuierer, I. Filipuzzi, S. Brachat, B. Bhullar, U. Plikat, Y. Abraham, M. Altorfer, T. Aust, L. Baeriswyl, R. Cerino, L. Chang, D. Estoppey, J. Eichenberger, M. Frederiksen, N. Hartmann, A. Hohendahl, B. Knapp, P. Krastel, N. Melin, F. Nigsch, E.J. Oakeley, V. Petitjean, F. Petersen, R. Riedl, E.K. Schmitt, F. Staedtler, C. Studer, J.A. Tallarico, S. Wetzel, M.C. Fishman, J.A. Porter, and N.R. Movva. 2014. High-resolution chemical dissection of a model eukaryote reveals targets, pathways and gene functions. Microbiological Research. 169:107–120. doi:10.1016/J.MICRES.2013.11.004. Hoffmann, P.C., T.A.M. Bharat, M.R. Wozny, J. Boulanger, E.A. Miller, and W. Kukulski. 2019. Tricalbins Contribute to Cellular Lipid Flux and Form Curved ER-PM Contacts that Are Bridged by Rod-Shaped Structures. Developmental Cell. 51:488. doi:10.1016/J.DEVCEL.2019.09.019. Hönscher, C., M. Mari, K. Auffarth, M. Bohnert, J. Griffith, W. Geerts, M. Van Der Laan, M. Cabrera, F. Reggiori, and C. Ungermann. 2014. Cellular Metabolism Regulates Contact Sites between Vacuoles and Mitochondria. Developmental Cell. 30:86–94. doi:10.1016/j.devcel.2014.06.006. Horenkamp, F.A., D.P. Valverde, J. Nunnari, and K.M. Reinisch. 2018. Molecular basis for sterol transport by StART‐like lipid transfer domains. EMBO J. 37:e98002. doi:10.15252/embj.201798002. Hsu, T.H., R.H. Chen, Y.H. Cheng, C.W. Wang, and H. Riezman. 2017. Lipid droplets are central organelles for meiosis II progression during yeast sporulation. Molecular Biology of the Cell. 28:440–451. doi:10.1091/mbc.E16-06-0375. 164 Huh, W.K., J. V Falvo, L.C. Gerke, A.S. Carroll, R.W. Howson, J.S. Weissman, and E.K. O’Shea. 2003. Global analysis of protein localization in budding yeast. Nature. 425:686–691. Huttlin, E.L., L. Ting, J. Raphael, J.A. Paulo, J. Wade, S.P. Gygi, E.L. Huttlin, L. Ting, R.J. Bruckner, F. Gebreab, M.P. Gygi, J. Szpyt, S. Tam, G. Zarraga, G. Colby, K. Baltier, R. Dong, V. Guarani, L.P. Vaites, A. Ordureau, J. Chick, B. Zhai, D. Kolippakkam, and J. Mintseris. 2015. The BioPlex Network : A Systematic Exploration of the Human Interactome. Cell. 162:425–440. doi:10.1016/j.cell.2015.06.043. Im, Y.J., S. Raychaudhuri, W.A. Prinz, and J.H. Hurley. 2005. Structural mechanism for sterol sensing and transport by OSBP-related proteins. Nature. 437:154–158. doi:10.1038/nature03923. Jain, A., and J.C.M. Holthuis. 2017. Membrane contact sites, ancient and central hubs of cellular lipid logistics. Biochimica et Biophysica Acta - Molecular Cell Research. 1864:1450–1458. doi:10.1016/j.bbamcr.2017.05.017. Janke, C., M.M. Magiera, N. Rathfelder, C. Taxis, S. Reber, H. Maekawa, A. Moreno-Borchart, G. Doenges, E. Schwob, E. Schiebel, and M. Knop. 2004. A versatile toolbox for PCR-based tagging of yeast genes: new fluorescent proteins, more markers and promoter substitution cassettes. Yeast. 21:947–962. doi:10.1002/yea.1142. Jansen, I.E., H. Ye, S. Heetveld, M.C. Lechler, H. Michels, R.I. Seinstra, S.J. Lubbe, V. Drouet, S. Lesage, E. Majounie, J.R. Gibbs, M.A. Nalls, M. Ryten, J.A. Botia, J. Vandrovcova, J. Simon-Sanchez, M. Castillo-Lizardo, P. Rizzu, C. Blauwendraat, A.K. Chouhan, Y. Li, P. Yogi, N. Amin, C.M. van Duijn, M.A. Nalls, V. Plagnol, D.G. Hernandez, M. Sharma, U.-M. Sheerin, M. Saad, J. SimónSánchez, C. Schulte, S. Lesage, S. Sveinbjörnsdóttir, S. Arepalli, R. Barker, Y. Ben-Shlomo, H.W. Berendse, D. Berg, K. Bhatia, R.M.A. de Bie, A. Biffi, B. Bloem, Z. Bochdanovits, M. Bonin, J.M. Bras, K. Brockmann, J. Brooks, D.J. Burn, E. Majounie, S. Lubbe, I.E. Jansen, R. Price, A. Nicolas, G. Charlesworth, C. Lungu, H. Chen, P.F. Chinnery, S. Chong, C.E. Clarke, M.R. Cookson, J.M. Cooper, J.C. Corvol, C. Counsell, P. Damier, J.-F. Dartigues, P. Deloukas, G. Deuschl, D.T. Dexter, K.D. van Dijk, A. Dillman, F. Durif, A. Dürr, S. Edkins, J.R. Evans, T. Foltynie, J. Dong, M. Gardner, J.R. Gibbs, A. Goate, E. Gray, R. Guerreiro, C. Harris, J.J. van Hilten, A. Hofman, A. Hollenbeck, J. Holton, M. Hu, I. Wurster, W. Mätzler, G. Hudson, S.E. Hunt, J. Huttenlocher, T. Illig, P.V. Jónsson, J.-C. Lambert, C. Langford, A. Lees, et al. 2017. Discovery and functional prioritization of Parkinson’s disease candidate genes from large-scale whole exome sequencing. Genome Biology. 18:22. doi:10.1186/s13059-017-1147-9. Jelokhani-Niaraki, M. 2022. Membrane Proteins: Structure, Function and Motion. Int J Mol Sci. 24:468. doi:10.3390/ijms24010468. Jentsch, J.A., I. Kiburu, K. Pandey, M. Timme, T. Ramlall, B. Levkau, J. Wu, D. Eliezer, O. Boudker, and A.K. Menon. 2018. Structural basis of sterol binding and transport by a 165 yeast StARkin domain. Journal of Biological Chemistry. 293:5522–5531. doi:10.1074/jbc.RA118.001881. Jeong, E., J. Kim, M. Go, and S.Y. Lee. 2020. Early estrogen‐induced gene 1 facilitates osteoclast formation through the inhibition of interferon regulatory factor 8 expression. The FASEB Journal. 34:12894–12906. doi:10.1096/fj.202001197R. Jeong, H., J. Park, H.-I. Kim, M. Lee, Y.-J. Ko, S. Lee, Y. Jun, and C. Lee. 2017. Mechanistic insight into the nucleus–vacuole junction based on the Vac8p–Nvj1p crystal structure. Proceedings of the National Academy of Sciences. 114:E4539–E4548. doi:10.1073/pnas.1701030114. Jesch, S.A., X. Zhao, M.T. Wells, and S.A. Henry. 2005. Genome-wide Analysis Reveals Inositol, Not Choline, as the Major Effector of Ino2p-Ino4p and Unfolded Protein Response Target Gene Expression in Yeast. Journal of Biological Chemistry. 280:9106–9118. doi:10.1074/jbc.M411770200. Jeyasimman, D., and Y. Saheki. 2020. SMP domain proteins in membrane lipid dynamics. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1865:158447. doi:10.1016/j.bbalip.2019.04.007. John Peter, A.T., B. Herrmann, D. Antunes, D. Rapaport, K.S. Dimmer, and B. Kornmann. 2017. Vps13-Mcp1 interact at vacuole-mitochondria interfaces and bypass ER-mitochondria contact sites. Journal of Cell Biology. 216:3219–3229. doi:10.1083/jcb.201610055. John Peter, A.T., S.N.S. Schie, N.J. Cheung, A.H. Michel, M. Peter, and B. Kornmann. 2022. Rewiring phospholipid biosynthesis reveals resilience to membrane perturbations and uncovers regulators of lipid homeostasis. The EMBO Journal. 41. doi:10.15252/embj.2021109998. Jumper, J., R. Evans, A. Pritzel, T. Green, M. Figurnov, O. Ronneberger, K. Tunyasuvunakool, R. Bates, A. Žídek, A. Potapenko, A. Bridgland, C. Meyer, S.A.A. Kohl, A.J. Ballard, A. Cowie, B. Romera-Paredes, S. Nikolov, R. Jain, J. Adler, T. Back, S. Petersen, D. Reiman, E. Clancy, M. Zielinski, M. Steinegger, M. Pacholska, T. Berghammer, S. Bodenstein, D. Silver, O. Vinyals, A.W. Senior, K. Kavukcuoglu, P. Kohli, and D. Hassabis. 2021. Highly accurate protein structure prediction with AlphaFold. Nature 2021 596:7873. 596:583–589. doi:10.1038/s41586-021-03819-2. Kaminska, J., W. Rzepnikowska, A. Polak, K. Flis, P. Soczewka, K. Bala, M. Sienko, M. Grynberg, P. Kaliszewski, A. Urbanek, K. Ayscough, and T. Zoladek. 2016. Phosphatidylinositol-3-phosphate regulates response of cells to proteotoxic stress. International Journal of Biochemistry and Cell Biology. 79:494–504. doi:10.1016/j.biocel.2016.08.007. 166 Kamiński, D.M. 2014. Recent progress in the study of the interactions of amphotericin B with cholesterol and ergosterol in lipid environments. Eur Biophys J. 43:453–467. doi:10.1007/s00249-014-0983-8. Kannan, M., S. Lahiri, L.K. Liu, V. Choudhary, and W.A. Prinz. 2017. Phosphatidylserine synthesis at membrane contact sites promotes its transport out of the ER. Journal of Lipid Research. 58:553–562. doi:10.1194/jlr.M072959. van der Kant, R., and J. Neefjes. 2014. Small regulators, major consequences - Ca2+ and cholesterol at the endosome-ER interface. J Cell Sci. 127:929–938. doi:10.1242/jcs.137539. Kaplan, M.R., and R.D. Simoni. 1985. Intracellular transport of phosphatidylcholine to the plasma membrane. J Cell Biol. 101:441–445. Kawano, S., Y. Tamura, R. Kojima, S. Bala, E. Asai, A.H. Michel, B. Kornmann, I. Riezman, H. Riezman, Y. Sakae, Y. Okamoto, and T. Endo. 2018. Structure-function insights into direct lipid transfer between membranes by Mmm 1-Mdm 12 of ERMES. Journal of Cell Biology. 217:959–974. doi:10.1083/jcb.201704119. Kay, J.G., and G.D. Fairn. 2019. Distribution, dynamics and functional roles of phosphatidylserine within the cell. Cell Communication and Signaling. 17:126. doi:10.1186/s12964-019-0438-z. Kay, J.G., and S. Grinstein. 2013. Phosphatidylserine-mediated cellular signaling. Adv Exp Med Biol. 991:177–193. doi:10.1007/978-94-007-6331-9_10. Kelley, M., J. Yochem, M. Krieg, A. Calixto, M.G. Heiman, A. Kuzmanov, V. Meli, M. Chalfie, M.B. Goodman, S. Shaham, A. Frand, and D.S. Fay. 2015. FBN-1, a fibrillin-related protein, is required for resistance of the epidermis to mechanical deformation during C. elegans embryogenesis. eLife. 23:e06565. doi:10.7554/eLife.06565. Kim, S., R. Coukos, F. Gao, and D. Krainc. 2022. Dysregulation of organelle membrane contact sites in neurological diseases. Neuron. 110:2386–2408. doi:10.1016/j.neuron.2022.04.020. Kishimoto, T., T. Mioka, E. Itoh, D.E. Williams, R.J. Andersen, and K. Tanaka. 2021. Phospholipid flippases and Sfk1 are essential for the retention of ergosterol in the plasma membrane. MBoC. 32:1374–1392. doi:10.1091/mbc.E20-11-0699. Koag, M.C., Y. Cheun, Y. Kou, H. Ouzon-Shubeita, K. Min, A.F. Monzingo, and S. Lee. 2013. Synthesis and structure of 16,22-diketocholesterol bound to oxysterol-binding protein Osh4. Steroids. 78:938–944. doi:10.1016/j.steroids.2013.05.016. Koegl, M., and P. Uetz. 2007. Improving yeast two-hybrid screening systems. Briefings in Functional Genomics. 6:302–312. doi:10.1093/bfgp/elm035. 167 Koharudin, L.M.I., W. Furey, H. Liu, Y.-J. Liu, and A.M. Gronenborn. 2009. The phox domain of sorting nexin 5 lacks phosphatidylinositol 3-phosphate (PtdIns(3)P) specificity and preferentially binds to phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2). J Biol Chem. 284:23697–23707. doi:10.1074/jbc.M109.008995. Koike, S., and R. Jahn. 2019. SNAREs define targeting specificity of trafficking vesicles by combinatorial interaction with tethering factors. Nature Communications. 10:1608. doi:10.1038/s41467-019-09617-9. Koizumi, K., and K.L. Gallagher. 2013. Identification of SHRUBBY , a SHORT-ROOT and SCARECROW interacting protein that controls root growth and radial patterning. Development. 140:1292–1300. doi:10.1242/dev.090761. Kolakowski, D., J. Kaminska, and T. Zoladek. 2020. The binding of the APT1 domains to phosphoinositides is regulated by metal ions in vitro. Biochimica et Biophysica Acta - Biomembranes. 1862:183349. doi:10.1016/j.bbamem.2020.183349. Kolakowski, D., W. Rzepnikowska, A. Kaniak-Golik, T. Zoladek, and J. Kaminska. 2021. The GTPase Arf1 Is a Determinant of Yeast Vps13 Localization to the Golgi Apparatus. Int J Mol Sci. 22:12274. doi:10.3390/ijms222212274. Kolehmainen, J., G.C.M. Black, A. Saarinen, K. Chandler, J. Clayton-Smith, A.-L. Träskelin, R. Perveen, S. Kivitie-Kallio, R. Norio, M. Warburg, J.-P. Fryns, A. de la Chapelle, and A.-E. Lehesjoki. 2003. Cohen syndrome is caused by mutations in a novel gene, COH1, encoding a transmembrane protein with a presumed role in vesicle-mediated sorting and intracellular protein transport. American Journal of Human Genetics. 72:1359–69. Kolehmainen, J., R. Wilkinson, A.-E. Lehesjoki, K. Chandler, S. Kivitie-Kallio, J. Clayton-Smith, A.-L. Träskelin, L. Waris, A. Saarinen, J. Khan, V. Gross-Tsur, E.I. Traboulsi, M. Warburg, J.-P. Fryns, R. Norio, G.C.M. Black, and F.D.C. Manson. 2004. Delineation of Cohen syndrome following a large-scale genotype-phenotype screen. American Journal of Human Genetics. 75:122–7. doi:10.1086/422197. Kopec, J., and G. Schneider. 2011. Comparison of fluorescence and light scattering based methods to assess formation and stability of protein–protein complexes. Journal of Structural Biology. 175:216–223. doi:10.1016/j.jsb.2011.04.006. Kopec, K.O., V. Alva, and A.N. Lupas. 2010. Homology of SMP domains to the TULIP superfamily of lipid-binding proteins provides a structural basis for lipid exchange between ER and mitochondria. Bioinformatics. 26:1927–1931. doi:10.1093/bioinformatics/btq326. Kornmann, B., E. Currie, S.R. Collins, M. Schuldiner, J. Nunnari, J.S. Weissman, and P. Walter. 2009. An ER-mitochondria tethering complex revealed by a synthetic biology screen. Science. 325:477–481. doi:10.1126/science.1175088. 168 Kors, S., J.L. Costello, and M. Schrader. 2022. VAP Proteins – From Organelle Tethers to Pathogenic Host Interactors and Their Role in Neuronal Disease. Front Cell Dev Biol. 10:895856. doi:10.3389/fcell.2022.895856. Kotani, T., H. Kirisako, M. Koizumi, Y. Ohsumi, and H. Nakatogawa. 2018a. The Atg2-Atg18 complex tethers pre-autophagosomal membranes to the endoplasmic reticulum for autophagosome formation. Proceedings of the National Academy of Sciences. 115:10363–10368. doi:10.1073/pnas.1806727115. Kotani, T., H. Kirisako, M. Koizumi, Y. Ohsumi, and H. Nakatogawa. 2018b. The Atg2-Atg18 complex tethers pre-autophagosomal membranes to the endoplasmic reticulum for autophagosome formation. Proceedings of the National Academy of Sciences of the United States of America. 115:10363–10368. doi:10.1073/pnas.1806727115. Krols, M., G. van Isterdael, B. Asselbergh, A. Kremer, S. Lippens, V. Timmerman, and S. Janssens. 2016. Mitochondria-associated membranes as hubs for neurodegeneration. Acta Neuropathologica. 131:505–23. doi:10.1007/s00401-015-1528-7. Kumar, N., M. Leonzino, W. Hancock-Cerutti, F.A. Horenkamp, P. Li, J.A. Lees, H. Wheeler, K.M. Reinisch, and P. De Camilli. 2018. VPS13A and VPS13C are lipid transport proteins differentially localized at ER contact sites. The Journal of Cell Biology. 217:3625–3639. doi:10.1083/jcb.201807019. Kvam, E., and D.S. Goldfarb. 2007. Nucleus-Vacuole Junctions and Piecemeal Microautophagy of the Nucleus in S. cerevisiae. Autophagy. 3:85–92. doi:10.4161/auto.3586. Lackner, L.L. 2019. The Expanding and Unexpected Functions of Mitochondria Contact Sites. Trends in Cell Biology. 29:580–590. doi:10.1016/j.tcb.2019.02.009. Lang, A.B., A.T. John Peter, P. Walter, and B. Kornmann. 2015. ER-mitochondrial junctions can be bypassed by dominant mutations in the endosomal protein Vps13. Journal of Cell Biology. 210:883–890. doi:10.1083/jcb.201502105. de Las Rivas, M., E. Lira-Navarrete, T.A. Gerken, and R. Hurtado-Guerrero. 2019. Polypeptide GalNAc-Ts: from redundancy to specificity. Curr Opin Struct Biol. 56:87–96. doi:10.1016/j.sbi.2018.12.007. Latham, C.F., S.L. Osborne, M.J. Cryle, and F.A. Meunier. 2006. Arachidonic acid potentiates exocytosis and allows neuronal SNARE complex to interact with Munc18a. J Neurochem. 0:1543–1554. doi:10.1111/j.1471-4159.2006.04286.x. Lesage, S., V. Drouet, E. Majounie, V. Deramecourt, M. Jacoupy, A. Nicolas, F. Cormier-Dequaire, S.M. Hassoun, C. Pujol, S. Ciura, Z. Erpapazoglou, T. Usenko, C.-A. Maurage, M. Sahbatou, S. Liebau, J. Ding, B. Bilgic, M. Emre, N. Erginel-Unaltuna, G. Guven, F. Tison, C. Tranchant, M. Vidailhet, J.-C. Corvol, P. Krack, A.-L. Leutenegger, M.A. Nalls, D.G. Hernandez, P. Heutink, J.R. Gibbs, J. Hardy, N.W. Wood, T. Gasser, A. 169 Durr, J.-F. Deleuze, M. Tzair, A. Destée, E. Lohmann, E. Kabashi, A. Singleton, O. Corti, A. Brice, French Parkinson’s Disease Genetics Study (PDG), and The International Parkinson’s Disease Genomics Consortium (IPDGC). 2016. Loss of VPS13C function in autosomal-recessive parkinsonism causes mitochondrial dysfunction and increases PINK1 / Parkin-dependent mitophagy. The American Journal of Human Genetics. 98:500–513. doi:10.1016/j.ajhg.2016.01.014. Lev, S. 2010. Non-vesicular lipid transport by lipid-transfer proteins and beyond. Nat Rev Mol Cell Biol. 11:739–750. doi:10.1038/nrm2971. Leventis, P.A., and S. Grinstein. 2010. The distribution and function of phosphatidylserine in cellular membranes. Annu Rev Biophys. 39:407–427. doi:10.1146/annurev.biophys.093008.131234. Levine, T. 2004. Short-range intracellular trafficking of small molecules across endoplasmic reticulum junctions. Trends in Cell Biology. 14:483–490. doi:10.1016/j.tcb.2004.07.017. Levine, T.P. 2022. Sequence Analysis and Structural Predictions of Lipid Transfer Bridges in the Repeating Beta Groove (RBG) Superfamily Reveal Past and Present Domain Variations Affecting Form, Function and Interactions of VPS13, ATG2, SHIP164, Hobbit and Tweek. Contact. 5:251525642211343. doi:10.1177/25152564221134328. Li, P.Q., J.A. Lees, C.P. Lusk, and K.M. Reinisch. 2020. Cryo-EM reconstruction of a VPS13 fragment reveals a long groove to channel lipids between membranes. The Journal of Cell Biology. 219. doi:10.1083/jcb.202001161. Li, X., J. Wang, E. Coutavas, H. Shi, Q. Hao, and G. Blobel. 2016. Structure of human Niemann-Pick C1 protein. Proceedings of the National Academy of Sciences. 113:8212–8217. doi:10.1073/pnas.1607795113. Lingwood, D., and K. Simons. 2010. Lipid Rafts As a Membrane-Organizing Principle. Science. 327:46–50. doi:10.1126/science.1174621. Liscum, L., and N.J. Munn. 1999. Intracellular cholesterol transport. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1438:19–37. doi:10.1016/S1388-1981(99)00043-8. Liu, L.K., V. Choudhary, A. Toulmay, and W.A. Prinz. 2016. An inducible ER–Golgi tether facilitates ceramide transport to alleviate lipotoxicity. Journal of Cell Biology. 1–17. Liu, T., X.-Y. Zhang, X.-H. He, J.-S. Geng, Y. Liu, D.-J. Kong, Q.-Y. Shi, F. Liu, W. Wei, and D. Pang. 2014. High Levels of BCOX1 Expression Are Associated with Poor Prognosis in Patients with Invasive Ductal Carcinomas of the Breast. PLoS ONE. 9:e86952. doi:10.1371/journal.pone.0086952. Loewen, C.J.R., and T.P. Levine. 2005. A highly conserved binding site in vesicle-associated membrane protein-associated protein (VAP) for the FFAT motif of lipid-binding 170 proteins. Journal of Biological Chemistry. 280:14097–14104. doi:10.1074/jbc.M500147200. Longtine, M.S., A. Mckenzie III, D.J. Demarini, N.G. Shah, A. Wach, A. Brachat, P. Philippsen, and J.R. Pringle. 1998. Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast. 14:953–961. doi:10.1002/(SICI)1097-0061(199807)14:10<953::AID-YEA293>3.0.CO;2-U. Lorent, J., K. Levental, L. Ganesan, G. Rivera-Longsworth, E. Sezgin, M. Doktorova, E. Lyman, and I. Levental. 2020. Plasma membranes are asymmetric in lipid unsaturation, packing, and protein shape. Nat Chem Biol. 16:644–652. doi:10.1038/s41589-020-0529-6. Lupo, F., E. Tibaldi, A. Matte, A.K. Sharma, A.M. Brunati, S.L. Alper, C. Zancanaro, D. Benati, A. Siciliano, M. Bertoldi, F. Zonta, A. Storch, R.H. Walker, A. Danek, B. Bader, A. Hermann, and L. De Franceschi. 2016. A new molecular link between defective autophagy and erythroid abnormalities in chorea-acanthocytosis. Blood. 128:2976–2987. doi:10.1182/blood-2016-07-727321. Maeda, S., C. Otomo, and T. Otomo. 2019. The autophagic membrane tether ATG2A transfers lipids between membranes. eLife. 8:e45777. doi:10.7554/eLife.45777. Maeda, S., H. Yamamoto, L.N. Kinch, C.M. Garza, S. Takahashi, C. Otomo, N.V. Grishin, S. Forli, N. Mizushima, and T. Otomo. 2020. Structure, lipid scrambling activity and role in autophagosome formation of ATG9A. Nat Struct Mol Biol. 27:1194–1201. doi:10.1038/s41594-020-00520-2. Maekawa, M., and G.D. Fairn. 2015. Complementary probes reveal that phosphatidylserine is required for the proper transbilayer distribution of cholesterol. Journal of Cell Science. 128:1422–1433. doi:10.1242/jcs.164715. Majounie, E., V. Deramecourt, S. Lesage, M. Jacoupy, A. Nicolas, F. Cormier-dequaire, S.M. Hassoun, C. Pujol, S. Ciura, Z. Erpapazoglou, T. Usenko, C. Maurage, M. Sahbatou, S. Liebau, J. Ding, C. Tranchant, M. Vidailhet, J. Corvol, A. Leutenegger, M.A. Nalls, D.G. Hernandez, P. Heutink, A. Brice, A. Singleton, and O. Corti. 2016. Loss of VPS13C Function in Autosomal-Recessive Parkinsonism Causes Mitochondrial Dysfunction and Increases PINK1 / Parkin-Dependent Mitophagy. American Journal of Human Genetics. 98:500–513. doi:10.1016/j.ajhg.2016.01.014. Manik, M.K., H. Yang, J. Tong, and Y.J. Im. 2017. Structure of Yeast OSBP-Related Protein Osh1 Reveals Key Determinants for Lipid Transport and Protein Targeting at the Nucleus-Vacuole Junction. Structure. 25:617-629.e3. doi:10.1016/j.str.2017.02.010. Marek, M., V. Vincenzetti, and S.G. Martin. 2020. Sterol biosensor reveals LAM-family Ltc1-dependent sterol flow to endosomes upon Arp2/3 inhibition. Journal of Cell Biology. 219:e202001147. doi:10.1083/jcb.202001147. 171 Marsh, D. 2007. Lateral Pressure Profile, Spontaneous Curvature Frustration, and the Incorporation and Conformation of Proteins in Membranes. Biophysical Journal. 93:3884–3899. doi:10.1529/biophysj.107.107938. Matoba, K., T. Kotani, A. Tsutsumi, T. Tsuji, T. Mori, D. Noshiro, Y. Sugita, N. Nomura, S. Iwata, Y. Ohsumi, T. Fujimoto, H. Nakatogawa, M. Kikkawa, and N.N. Noda. 2020. Atg9 is a lipid scramblase that mediates autophagosomal membrane expansion. Nature Structural & Molecular Biology. 1–9. doi:10.1038/s41594-020-00518-w. van Meer, G. 2005. Cellular lipidomics. EMBO J. 24:3159–3165. doi:10.1038/sj.emboj.7600798. van Meer, G. 2011. Dynamic Transbilayer Lipid Asymmetry. Cold Spring Harb Perspect Biol. 3:a004671. doi:10.1101/cshperspect.a004671. van Meer, G., D.R. Voelker, and G.W. Feigenson. 2008. Membrane lipids: Where they are and how they behave. Nature Reviews Molecular Cell Biology. 9:112–124. doi:10.1038/nrm2330. Mehla, J., J.H. Caufield, N. Sakhawalkar, and P. Uetz. 2017. A comparison of two hybrid approaches for detecting protein-protein interactions. Methods Enzymol. 586:333–358. doi:10.1016/bs.mie.2016.10.020. Mesmin, B., B. Antonny, and G. Drin. 2013. Insights into the mechanisms of sterol transport between organelles. Cell. Mol. Life Sci. 70:3405–3421. doi:10.1007/s00018-012-1247-3. Mesmin, B., and F.R. Maxfield. 2009. Intracellular Sterol Dynamics. Biochim Biophys Acta. 1791:636–645. doi:10.1016/j.bbalip.2009.03.002. Meunier, F.A., S.L. Osborne, G.R.V. Hammond, F.T. Cooke, P.J. Parker, J. Domin, and G. Schiavo. 2005. Phosphatidylinositol 3-Kinase C2α Is Essential for ATP-dependent Priming of Neurosecretory Granule Exocytosis. MBoC. 16:4841–4851. doi:10.1091/mbc.e05-02-0171. Mi, H., A. Muruganujan, D. Ebert, X. Huang, and P.D. Thomas. 2019. PANTHER version 14: more genomes, a new PANTHER GO-slim and improvements in enrichment analysis tools. Nucleic Acids Research. 47:D419–D426. doi:10.1093/nar/gky1038. Michnick, S.W., P.H. Ear, C. Landry, M.K. Malleshaiah, and V. Messier. 2010. Chapter 14 - A Toolkit of Protein-Fragment Complementation Assays for Studying and Dissecting Large-Scale and Dynamic Protein–Protein Interactions in Living Cells. In Methods in Enzymology. Academic Press. 335–368. Mikitova, V., and T.P. Levine. 2012. Analysis of the key elements of FFAT-like motifs identifies new proteins that potentially bind VAP on the ER, including two AKAPs and FAPP2. PLoS ONE. 7:e30455. doi:10.1371/journal.pone.0030455. 172 Miliara, X., T. Tatsuta, J.-L. Berry, S.L. Rouse, K. Solak, D.S. Chorev, D. Wu, C.V. Robinson, S. Matthews, and T. Langer. 2019. Structural determinants of lipid specificity within Ups/PRELI lipid transfer proteins. Nat Commun. 10:1130. doi:10.1038/s41467-019-09089-x. Milosevic, I. 2005. Plasmalemmal Phosphatidylinositol-4,5-Bisphosphate Level Regulates the Releasable Vesicle Pool Size in Chromaffin Cells. Journal of Neuroscience. 25:2557–2565. doi:10.1523/JNEUROSCI.3761-04.2005. Min, S.-W., W.-P. Chang, and T.C. Südhof. 2007. E-Syts, a family of membranous Ca2+-sensor proteins with multiple C2 domains. Proceedings of the National Academy of Sciences. 104:3823–3828. doi:10.1073/pnas.0611725104. Mirdita, M., K. Schütze, Y. Moriwaki, L. Heo, S. Ovchinnikov, and M. Steinegger. 2022. ColabFold: making protein folding accessible to all. Nat Methods. 19:679–682. doi:10.1038/s41592-022-01488-1. Mironov, A.A., A. Mironov, J. Derganc, and G.V. Beznoussenko. 2020. Membrane Curvature, Trans-Membrane Area Asymmetry, Budding, Fission and Organelle Geometry. Int J Mol Sci. 21:7594. doi:10.3390/ijms21207594. Mollinedo, F., and C. Gajate. 2020. Lipid rafts as signaling hubs in cancer cell survival/death and invasion: implications in tumor progression and therapy. J Lipid Res. 61:611–635. doi:10.1194/jlr.TR119000439. Montigny, C., J. Lyons, P. Champeil, P. Nissen, and G. Lenoir. 2016. On the molecular mechanism of flippase- and scramblase-mediated phospholipid transport. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1861:767–783. doi:10.1016/j.bbalip.2015.12.020. Moser von Filseck, J., A. Čopič, V. Delfosse, S. Vanni, C.L. Jackson, W. Bourguet, and G. Drin. 2015. Phosphatidylserine transport by ORP/Osh proteins is driven by phosphatidylinositol 4-phosphate. Science. 349:432–436. doi:10.1126/science.aab1346. Muñoz-Braceras, S., R. Calvo, and R. Escalante. 2015. TipC and the chorea-acanthocytosis protein VPS13A regulate autophagy in Dictyostelium and human HeLa cells. Autophagy. 11:918–927. doi:10.1080/15548627.2015.1034413. Muñoz-Braceras, S., A.R. Tornero-Écija, O. Vincent, and R. Escalante. 2019. VPS13A is closely associated with mitochondria and is required for efficient lysosomal degradation. Disease Models & Mechanisms. 12. doi:10.1242/dmm.036681. Murley, A., and J. Nunnari. 2016. The Emerging Network of Mitochondria-Organelle Contacts. Molecular Cell. 61:648–653. doi:10.1016/j.molcel.2016.01.031. 173 Murley, A., R.D. Sarsam, A. Toulmay, J. Yamada, W.A. Prinz, and J. Nunnari. 2015. Ltc1 is an ER-localized sterol transporter and a component of ER–mitochondria and ER–vacuole contacts. Journal of Cell Biology. 209:539–548. doi:10.1083/jcb.201502033. Murphy, S.E., and T.P. Levine. 2016. VAP, a Versatile Access Point for the Endoplasmic Reticulum: Review and analysis of FFAT-like motifs in the VAPome. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids. 1861:952–961. doi:10.1016/j.bbalip.2016.02.009. Muthusamy, B.-P., P. Natarajan, X. Zhou, and T.R. Graham. 2009. Linking Phospholipid flippases to vesicle-mediated protein transport. Biochim Biophys Acta. 1791:612–619. doi:10.1016/j.bbalip.2009.03.004. Nakamura, T.S., Y. Suda, K. Muneshige, Y. Fujieda, Y. Okumura, I. Inoue, T. Tanaka, T. Takahashi, H. Nakanishi, X.-D. Gao, Y. Okada, A.M. Neiman, and H. Tachikawa. 2021. Suppression of Vps13 adaptor protein mutants reveals a central role for PI4P in regulating prospore membrane extension. PLoS Genet. 17:e1009727. doi:10.1371/journal.pgen.1009727. Nakatsu, F., and A. Kawasaki. 2021. Functions of Oxysterol-Binding Proteins at Membrane Contact Sites and Their Control by Phosphoinositide Metabolism. Frontiers in Cell and Developmental Biology. 9. Neuman, S.D., and A. Bashirullah. 2018. Hobbit regulates intracellular trafficking to drive insulin-dependent growth during Drosophila development. Development. 145. doi:10.1242/dev.161356. Neuman, S.D., J.R. Jorgensen, A.T. Cavanagh, J.T. Smyth, J.E. Selegue, S.D. Emr, and A. Bashirullah. 2022a. The Hob proteins are novel and conserved lipid-binding proteins at ER-PM contact sites. J Cell Sci. 135:jcs259086. doi:10.1242/jcs.259086. Neuman, S.D., T.P. Levine, and A. Bashirullah. 2022b. A novel superfamily of bridge-like lipid transfer proteins. Trends in Cell Biology. 32:962–974. doi:10.1016/j.tcb.2022.03.011. Nishimura, A.L., M. Mitne-Neto, H.C.A. Silva, A. Richieri-Costa, S. Middleton, D. Cascio, F. Kok, J.R.M. Oliveira, T. Gillingwater, J. Webb, P. Skehel, and M. Zatz. 2004. A mutation in the vesicle-trafficking protein VAPB causes late-onset spinal muscular atrophy and amyotrophic lateral sclerosis. American Journal of Human Genetics. 75:822–831. doi:10.1086/425287. Olkkonen, V.M., and S. Li. 2013. Oxysterol-binding proteins: sterol and phosphoinositide sensors coordinating transport, signaling and metabolism. Prog Lipid Res. 52:529–538. doi:10.1016/j.plipres.2013.06.004. 174 Omnus, D.J., A. Cadou, F.B. Thomas, J.M. Bader, N. Soh, G.H.C. Chung, A.N. Vaughan, and C.J. Stefan. 2020. A heat-sensitive Osh protein controls PI4P polarity. BMC Biology. 18:28. doi:10.1186/s12915-020-0758-x. Osawa, T., T. Kotani, T. Kawaoka, E. Hirata, K. Suzuki, H. Nakatogawa, Y. Ohsumi, and N.N. Noda. 2019. Atg2 mediates direct lipid transfer between membranes for autophagosome formation. Nature Structural & Molecular Biology. 26:281–288. doi:10.1038/s41594-019-0203-4. Paillusson, S., R. Stoica, P. Gomez-Suaga, D.H.W. Lau, S. Mueller, T. Miller, and C.C.J. Miller. 2016. There’s Something Wrong with my MAM; the ER-Mitochondria Axis and Neurodegenerative Diseases. Trends in Neurosciences. 39:146–157. doi:10.1016/j.tins.2016.01.008. Pan, X., P. Roberts, Y. Chen, E. Kvam, N. Shulga, K. Huang, S. Lemmon, and D.S. Goldfarb. 2000. Nucleus–Vacuole Junctions in Saccharomyces cerevisiae Are Formed Through the Direct Interaction of Vac8p with Nvj1p. MBoC. 11:2445–2457. doi:10.1091/mbc.11.7.2445. Park, J.-S., and A.M. Neiman. 2012. VPS13 regulates membrane morphogenesis during sporulation in Saccharomyces cerevisiae. Journal of Cell Science. 125:3004–11. doi:10.1242/jcs.105114. Park, J.S., and A.M. Neiman. 2020. XK is a partner for VPS13A: a molecular link between Chorea-Acanthocytosis and McLeod Syndrome. Molecular Biology of the Cell. 31:2425–2436. doi:10.1091/MBC.E19-08-0439-T. Park, J.-S., Y. Okumura, H. Tachikawa, and A.M. Neiman. 2013. SPO71 encodes a developmental stage-specific partner for Vps13 in Saccharomyces cerevisiae. Eukaryotic Cell. 12:1530–1537. doi:10.1128/EC.00239-13. Park, J.-S., M.K. Thorsness, R. Policastro, L.L. McGoldrick, N.M. Hollingsworth, P.E. Thorsness, and A.M. Neiman. 2016. Yeast Vps13 promotes mitochondrial function and is localized at membrane contact sites. Molecular Biology of the Cell. 27:2435–2449. doi:10.1091/mbc.E16-02-0112. Parsons, A.B., A. Lopez, I.E. Givoni, D.E. Williams, C.A. Gray, J. Porter, G. Chua, R. Sopko, R.L. Brost, C.-H. Ho, J. Wang, T. Ketela, C. Brenner, J.A. Brill, G.E. Fernandez, T.C. Lorenz, G.S. Payne, S. Ishihara, Y. Ohya, B. Andrews, T.R. Hughes, B.J. Frey, T.R. Graham, R.J. Andersen, and C. Boone. 2006. Exploring the Mode-of-Action of Bioactive Compounds by Chemical-Genetic Profiling in Yeast. Cell. 126:611–625. doi:10.1016/j.cell.2006.06.040. Pashkova, N., T.A. Peterson, V. Krishnamani, P. Breheny, M. Stamnes, and R.C. Piper. 2016. DEEPN as an Approach for Batch Processing of Yeast 2-Hybrid Interactions. Cell Reports. 17:303–315. doi:10.1016/j.celrep.2016.08.095. 175 Peretti, D., S. Kim, R. Tufi, and S. Lev. 2020. Lipid Transfer Proteins and Membrane Contact Sites in Human Cancer. Front Cell Dev Biol. 7:371. doi:10.3389/fcell.2019.00371. Pérez-Lara, Á., and R. Jahn. 2015. Extended synaptotagmins (E-Syts): Architecture and dynamics of membrane contact sites revealed. Proceedings of the National Academy of Sciences. 112:4837–4838. doi:10.1073/pnas.1504487112. Pettersen, E.F., T.D. Goddard, C.C. Huang, G.S. Couch, D.M. Greenblatt, E.C. Meng, and T.E. Ferrin. 2004. UCSF Chimera - A visualization system for exploratory research and analysis. J. Comput. Chem. 25:1605–1612. doi:10.1002/jcc.20084. Pettersen, E.F., T.D. Goddard, C.C. Huang, E.C. Meng, G.S. Couch, T.I. Croll, J.H. Morris, and T.E. Ferrin. 2021. UCSF ChimeraX: Structure visualization for researchers, educators, and developers. Protein Science. 30:70–82. doi:10.1002/pro.3943. Phillips, M.J., and G.K. Voeltz. 2016. Structure and function of ER membrane contact sites with other organelles. Nature Reviews Molecular Cell Biology. 17:69–82. doi:10.1038/nrm.2015.8. Pomorski, T., and A.K. Menon. 2006. Lipid flippases and their biological functions. Cell. Mol. Life Sci. 63:2908–2921. doi:10.1007/s00018-006-6167-7. Popa, S., J. Villeneuve, S. Stewart, E. Perez Garcia, A. Petrunkina Harrison, and K. Moreau. 2020. Genome-wide CRISPR screening identifies new regulators of glycoprotein secretion. Wellcome Open Res. 4:119. doi:10.12688/wellcomeopenres.15232.2. Prinz, W.A., A. Toulmay, and T. Balla. 2020. The functional universe of membrane contact sites. Nat Rev Mol Cell Biol. 21:7–24. doi:10.1038/s41580-019-0180-9. Rampoldi, L., C. Dobson-Stone, J.P. Rubio, A. Danek, R.M. Chalmers, N.W. Wood, C. Verellen, X. Ferrer, A. Malandrini, G.M. Fabrizi, R. Brown, J. Vance, M. Pericak-Vance, G. Rudolf, S. Carrè, E. Alonso, M. Manfredi, A.H. Németh, and A.P. Monaco. 2001. A conserved sorting-associated protein is mutant in chorea-acanthocytosis. Nature Genetics. 28:119–120. doi:10.1038/88821. Ramseyer, V.D., V.A. Kimler, and J.G. Granneman. 2017. Vacuolar protein sorting 13C is a novel lipid droplet protein that inhibits lipolysis in brown adipocytes. Molecular Metabolism. 1–14. doi:10.1016/j.molmet.2017.10.014. Ramseyer, V.D., V.A. Kimler, and J.G. Granneman. 2018. Vacuolar protein sorting 13C is a novel lipid droplet protein that inhibits lipolysis in brown adipocytes. Molecular Metabolism. 7:57–70. doi:10.1016/j.molmet.2017.10.014. Rauthan, M., and M. Pilon. 2011. The mevalonate pathway in C. elegans. Lipids Health Dis. 10:243. doi:10.1186/1476-511X-10-243. 176 Reinisch, K.M., and W.A. Prinz. 2021. Mechanisms of nonvesicular lipid transport. Journal of Cell Biology. 220:e202012058. doi:10.1083/jcb.202012058. Ripa, I., S. Andreu, J.A. López-Guerrero, and R. Bello-Morales. 2021. Membrane Rafts: Portals for Viral Entry. Frontiers in Microbiology. 12. Roberts, P., S. Moshitch-Moshkovitz, E. Kvam, E. O’Toole, M. Winey, and D.S. Goldfarb. 2003. Piecemeal microautophagy of nucleus in Saccharomyces cerevisiae. Molecular Biology of the Cell. 14:129–141. doi:10.1091/mbc.E02-08-0483. Roelants, F.M., N. Chauhan, A. Muir, J.C. Davis, A.K. Menon, T.P. Levine, and J. Thorner. 2018. TOR complex 2-regulated protein kinase Ypk1 controls sterol distribution by inhibiting StARkin domain-containing proteins located at plasma membrane-endoplasmic reticulum contact sites. Molecular Biology of the Cell. 29:2128–2136. doi:10.1091/mbc.E18-04-0229. Rogers, L.D., and L.J. Foster. 2007. The dynamic phagosomal proteome and the contribution of the endoplasmic reticulum. Proceedings of the National Academy of Sciences. 104:18520–18525. doi:10.1073/pnas.0705801104. Roulis, E., C. Hyland, R. Flower, C. Gassner, H.H. Jung, and B.M. Frey. 2018. Molecular Basis and Clinical Overview of McLeod Syndrome Compared with Other Neuroacanthocytosis Syndromes: A Review. JAMA Neurology. 75:1554–1562. doi:10.1001/jamaneurol.2018.2166. Ruiz, N., R.M. Davis, and S. Kumar. 2021. YhdP, TamB, and YdbH Are Redundant but Essential for Growth and Lipid Homeostasis of the Gram-Negative Outer Membrane. mBio. 12:e02714-21. doi:10.1128/mBio.02714-21. Rzepnikowska, W., K. Flis, J. Kaminska, M. Grynberg, A. Urbanek, K.R. Ayscough, and T. Zoladek. 2017. Amino acid substitution equivalent to human chorea-acanthocytosis I2771R in yeast Vps13 protein affects its binding to phosphatidylinositol 3-phosphate. Human Molecular Genetics. 26:1497–1510. doi:10.1093/hmg/ddx054. Saheki, Y., and P. De Camilli. 2017. The Extended-Synaptotagmins. Biochim Biophys Acta. 1864:1490–1493. doi:10.1016/j.bbamcr.2017.03.013. de Saint-Jean, M., V. Delfosse, D. Douguet, G. Chicanne, B. Payrastre, W. Bourguet, B. Antonny, and G. Drin. 2011. Osh4p exchanges sterols for phosphatidylinositol 4-phosphate between lipid bilayers. J Cell Biol. 195:965–978. doi:10.1083/jcb.201104062. Saleh, M.-C., R.P. van Rij, A. Hekele, A. Gillis, E. Foley, P.H. O’Farrell, and R. Andino. 2006. The endocytic pathway mediates cell entry of dsRNA to induce RNAi silencing. Nat Cell Biol. 8:793–802. doi:10.1038/ncb1439. 177 Samavarchi-Tehrani, P., R. Samson, and A.-C. Gingras. 2020. Proximity Dependent Biotinylation: Key Enzymes and Adaptation to Proteomics Approaches. Mol Cell Proteomics. 19:757–773. doi:10.1074/mcp.R120.001941. Schauder, C.M., X. Wu, Y. Saheki, P. Narayanaswamy, F. Torta, M.R. Wenk, P. De Camilli, and K.M. Reinisch. 2014. Structure of a lipid-bound Extended-Synaptotagmin indicates a role in lipid transfer. Nature. 510:552–555. doi:10.1038/nature13269. Schindelin, J., I. Arganda-Carreras, E. Frise, V. Kaynig, M. Longair, T. Pietzsch, S. Preibisch, C. Rueden, S. Saalfeld, B. Schmid, J.-Y. Tinevez, D.J. White, V. Hartenstein, K. Eliceiri, P. Tomancak, and A. Cardona. 2012. Fiji: an open-source platform for biological-image analysis. Nature Methods. 9:676–82. doi:10.1038/nmeth.2019. Schütter, M., P. Giavalisco, S. Brodesser, and M. Graef. 2020. Local Fatty Acid Channeling into Phospholipid Synthesis Drives Phagophore Expansion during Autophagy. Cell. 180:135-149.e14. doi:10.1016/j.cell.2019.12.005. Scorrano, L., M.A. De Matteis, S. Emr, F. Giordano, G. Hajnóczky, B. Kornmann, L.L. Lackner, T.P. Levine, L. Pellegrini, K. Reinisch, R. Rizzuto, T. Simmen, H. Stenmark, C. Ungermann, and M. Schuldiner. 2019. Coming together to define membrane contact sites. Nature Communications. 10:1287. doi:10.1038/s41467-019-09253-3. Sebastian, T.T., R.D. Baldridge, P. Xu, and T.R. Graham. 2012. Phospholipid flippases: building asymmetric membranes and transport vesicles. Biochim Biophys Acta. 1821:1068–1077. doi:10.1016/j.bbalip.2011.12.007. Seifert, W., M. Holder-Espinasse, S. Spranger, M. Hoeltzenbein, E. Rossier, H. Dollfus, D. Lacombe, A. Verloes, K.H. Chrzanowska, G.H.B. Maegawa, D. Chitayat, D. Kotzot, D. Huhle, P. Meinecke, B. Albrecht, I. Mathijssen, B. Leheup, K. Raile, H.C. Hennies, and D. Horn. 2006. Mutational spectrum of COH1 and clinical heterogeneity in Cohen syndrome. Journal of Medical Genetics. 43:e22. doi:10.1136/jmg.2005.039867. Seifert, W., J. Kühnisch, T. Maritzen, D. Horn, V. Haucke, and H.C. Hennies. 2011. Cohen syndrome-associated protein, COH1, is a novel, giant Golgi matrix protein required for Golgi integrity. Journal of Biological Chemistry. 286:37665–37675. doi:10.1074/jbc.M111.267971. Seifert, W., J. Kühnisch, T. Maritzen, S. Lommatzsch, H.C. Hennies, S. Bachmann, D. Horn, and V. Haucke. 2015. Cohen syndrome-associated protein COH1 physically and functionally interacts with the small GTPase RAB6 at the Golgi complex and directs neurite outgrowth. The Journal of Biological Chemistry. 290:3349–58. doi:10.1074/jbc.M114.608174. Senisterra, G.A., and P.J. Finerty. 2009. High throughput methods of assessing protein stability and aggregation. Mol Biosyst. 5:217–223. doi:10.1039/b814377c. 178 Senisterra, G.A., H. Ghanei, G. Khutoreskaya, E. Dobrovetsky, A.M. Edwards, G.G. Privé, and M. Vedadi. 2010. Assessing the Stability of Membrane Proteins to Detect Ligand Binding Using Differential Static Light Scattering. J Biomol Screen. 15:314–320. doi:10.1177/1087057109357117. Seong, E., R. Insolera, M. Dulovic, E.-J. Kamsteeg, J. Trinh, N. Brüggemann, E. Sandford, S. Li, A.B. Ozel, J.Z. Li, T. Jewett, A.J.A. Kievit, A. Münchau, V. Shakkottai, C. Klein, C.A. Collins, K. Lohmann, B.P. van de Warrenburg, and M. Burmeister. 2018. Mutations in VPS13D lead to a new recessive ataxia with spasticity and mitochondrial defects. Annals of Neurology. 83:1075–1088. doi:10.1002/ana.25220. Shannon, P., A. Markiel, O. Ozier, N.S. Baliga, J.T. Wang, D. Ramage, N. Amin, B. Schwikowski, and T. Ideker. 2003. Cytoscape: A Software Environment for Integrated Models of Biomolecular Interaction Networks. Genome Res. 13:2498–2504. doi:10.1101/gr.1239303. Sheetz, M.P., and S.J. Singer. 1974. Biological Membranes as Bilayer Couples. A Molecular Mechanism of Drug-Erythrocyte Interactions. Proc Natl Acad Sci U S A. 71:4457–4461. Shpilka, T., E. Welter, N. Borovsky, N. Amar, M. Mari, F. Reggiori, and Z. Elazar. 2015. Lipid droplets and their component triglycerides and steryl esters regulate autophagosome biogenesis. The EMBO Journal. 34:2117–2131. doi:10.15252/embj.201490315. Shui, W., L. Sheu, J. Liu, B. Smart, C.J. Petzold, T.Y. Hsieh, A. Pitcher, J.D. Keasling, and C.R. Bertozzi. 2008. Membrane proteomics of phagosomes suggests a connection to autophagy. Proceedings of the National Academy of Sciences. 105:16952–16957. doi:10.1073/pnas.0809218105. Singh, R.P., B.R. Brooks, and J.B. Klauda. 2009. Binding and Release of Cholesterol in the Osh4 Protein of Yeast. Proteins. 75:468–477. doi:10.1002/prot.22263. Söding, J., A. Biegert, and A.N. Lupas. 2005. The HHpred interactive server for protein homology detection and structure prediction. Nucleic Acids Research. 33:W244. doi:10.1093/nar/gki408. Solanko, L.M., D.P. Sullivan, Y.Y. Sere, M. Szomek, A. Lunding, K.A. Solanko, A. Pizovic, L.D. Stanchev, T.G. Pomorski, A.K. Menon, and D. Wüstner. 2018. Ergosterol is mainly located in the cytoplasmic leaflet of the yeast plasma membrane. Traffic. 19:198–214. doi:10.1111/tra.12545. Song, J., W. Yang, I.M. Shih, Z. Zhang, and J. Bai. 2006. Identification of BCOX1, a novel gene overexpressed in breast cancer. Biochimica et Biophysica Acta - General Subjects. 1760:62–69. doi:10.1016/j.bbagen.2005.09.017. Sperandeo, P., A.M. Martorana, and A. Polissi. 2017. The lipopolysaccharide transport (Lpt) machinery: A nonconventional transporter for lipopolysaccharide assembly at the outer 179 membrane of Gram-negative bacteria. J Biol Chem. 292:17981–17990. doi:10.1074/jbc.R117.802512. Sridhar, V. 2023. Investigating the role of the putative lipid transport protein Fmp27 at ER-PM membrane contact sites. University of British Columbia. Stefan, C.J., A. Audhya, and S.D. Emr. 2002. The Yeast Synaptojanin-like Proteins Control the Cellular Distribution of Phosphatidylinositol (4,5)-Bisphosphate. Mol Biol Cell. 13:542–557. doi:10.1091/mbc.01-10-0476. Stefan, C.J., W.S. Trimble, S. Grinstein, G. Drin, K. Reinisch, P. De Camilli, S. Cohen, A.M. Valm, J. Lippincott-Schwartz, T.P. Levine, D.B. Iaea, F.R. Maxfield, C.E. Futter, E.R. Eden, D. Judith, A.R. van Vliet, P. Agostinis, S.A. Tooze, A. Sugiura, and H.M. McBride. 2017. Membrane dynamics and organelle biogenesis—lipid pipelines and vesicular carriers. BMC Biol. 15:102, s12915-017-0432–0. doi:10.1186/s12915-017-0432-0. Tai, S.L., V.M. Boer, P. Daran-Lapujade, M.C. Walsh, J.H. de Winde, J.-M. Daran, and J.T. Pronk. 2005. Two-dimensional Transcriptome Analysis in Chemostat Cultures. Journal of Biological Chemistry. 280:437–447. doi:10.1074/jbc.M410573200. Tamura, N., T. Nishimura, Y. Sakamaki, I. Koyama-Honda, H. Yamamoto, and N. Mizushima. 2017. Differential requirement for ATG2A domains for localization to autophagic membranes and lipid droplets. FEBS Letters. 591:3819–3830. doi:10.1002/1873-3468.12901. Tamura, Y., O. Onguka, K. Itoh, T. Endo, M. Iijima, S.M. Claypool, and H. Sesaki. 2012. Phosphatidylethanolamine Biosynthesis in Mitochondria. J Biol Chem. 287:43961–43971. doi:10.1074/jbc.M112.390997. Tan, T., C. Ozbalci, B. Brugger, D. Rapaport, and K.S. Dimmer. 2013. Mcp1 and Mcp2 , two novel proteins involved in mitochondrial lipid homeostasis. Journal of Cell Science. 126:3563–3574. doi:10.1242/jcs.121244. Tanguy, E., O. Carmon, Q. Wang, L. Jeandel, S. Chasserot-Golaz, M. Montero-Hadjadje, and N. Vitale. 2016. Lipids implicated in the journey of a secretory granule: from biogenesis to fusion. J. Neurochem. 137:904–912. doi:10.1111/jnc.13577. The Gene Ontology Consortium, S. Carbon, E. Douglass, B.M. Good, D.R. Unni, N.L. Harris, C.J. Mungall, S. Basu, R.L. Chisholm, R.J. Dodson, E. Hartline, P. Fey, P.D. Thomas, L.-P. Albou, D. Ebert, M.J. Kesling, H. Mi, A. Muruganujan, X. Huang, T. Mushayahama, S.A. LaBonte, D.A. Siegele, G. Antonazzo, H. Attrill, N.H. Brown, P. Garapati, S.J. Marygold, V. Trovisco, G. dos Santos, K. Falls, C. Tabone, P. Zhou, J.L. Goodman, V.B. Strelets, J. Thurmond, P. Garmiri, R. Ishtiaq, M. Rodríguez-López, M.L. Acencio, M. Kuiper, A. Lægreid, C. Logie, R.C. Lovering, B. Kramarz, S.C.C. Saverimuttu, S.M. Pinheiro, H. Gunn, R. Su, K.E. Thurlow, M. Chibucos, M. Giglio, S. Nadendla, J. Munro, 180 R. Jackson, M.J. Duesbury, N. Del-Toro, B.H.M. Meldal, K. Paneerselvam, L. Perfetto, P. Porras, S. Orchard, A. Shrivastava, H.-Y. Chang, R.D. Finn, A.L. Mitchell, N.D. Rawlings, L. Richardson, A. Sangrador-Vegas, J.A. Blake, K.R. Christie, M.E. Dolan, H.J. Drabkin, D.P. Hill, L. Ni, D.M. Sitnikov, M.A. Harris, S.G. Oliver, K. Rutherford, V. Wood, J. Hayles, J. Bähler, E.R. Bolton, J.L. De Pons, M.R. Dwinell, G.T. Hayman, M.L. Kaldunski, A.E. Kwitek, S.J.F. Laulederkind, C. Plasterer, M.A. Tutaj, M. Vedi, S.-J. Wang, P. D’Eustachio, L. Matthews, J.P. Balhoff, S.A. Aleksander, M.J. Alexander, J.M. Cherry, et al. 2021. The Gene Ontology resource: enriching a GOld mine. Nucleic Acids Research. 49:D325–D334. doi:10.1093/nar/gkaa1113. Tomiyasu, A., M. Nakamura, M. Ichiba, S. Ueno, S. Saiki, M. Morimoto, J. Kobal, Y. Kageyama, T. Inui, K. Wakabayashi, T. Yamada, Y. Kanemori, H.H. Jung, H. Tanaka, S. Orimo, Z. Afawi, I. Blatt, J. Aasly, H. Ujike, D. Babovic-Vuksanovic, K.A. Josephs, R. Tohge, G.R. Rodrigues, N. Dupré, H. Yamada, F. Yokochi, K. Kotschet, T. Takei, M. Rudzińska, A. Szczudlik, S. Penco, M. Fujiwara, K. Tojo, and A. Sano. 2011. Novel pathogenic mutations and copy number variations in the VPS13A Gene in patients with chorea-acanthocytosis. American Journal of Medical Genetics Part B: Neuropsychiatric Genetics. 156:620–631. doi:10.1002/ajmg.b.31206. Tong, J., M.K. Manik, and Y.J. Im. 2018. Structural basis of sterol recognition and nonvesicular transport by lipid transfer proteins anchored at membrane contact sites. Proc. Natl. Acad. Sci. U.S.A. 115. doi:10.1073/pnas.1719709115. Tong, J., L. Tan, and Y.J. Im. 2021. Structure of human ORP3 ORD reveals conservation of a key function and ligand specificity in OSBP-related proteins. PLoS One. 16:e0248781. doi:10.1371/journal.pone.0248781. Tong, J., H. Yang, H. Yang, S.H. Eom, and Y.J. Im. 2013. Structure of Osh3 reveals a conserved mode of phosphoinositide binding in oxysterol-binding proteins. Structure. 21:1203–1213. doi:10.1016/j.str.2013.05.007. Topolska, M., F.M. Roelants, E.P. Si, and J. Thorner. 2020. TORC2-Dependent Ypk1-Mediated Phosphorylation of Lam2/Ltc4 Disrupts Its Association with the β-Propeller Protein Laf1 at Endoplasmic Reticulum-Plasma Membrane Contact Sites in the Yeast Saccharomyces cerevisiae. Biomolecules. 10:1598. doi:10.3390/biom10121598. Tornero-Écija, A., A. Zapata-Del-Baño, L. Antón-Esteban, O. Vincent, and R. Escalante. 2023. The association of lipid transfer protein VPS13A with endosomes is mediated by sorting nexin SNX5. Life Sci Alliance. 6:e202201852. doi:10.26508/lsa.202201852. Toulmay, A., F.B. Whittle, J. Yang, X. Bai, J. Diarra, S. Banerjee, T.P. Levine, A. Golden, and W.A. Prinz. 2022. Vps13-like proteins provide phosphatidylethanolamine for GPI anchor synthesis in the ER. Journal of Cell Biology. 221:e202111095. doi:10.1083/jcb.202111095. 181 Trost, M., L. English, S. Lemieux, M. Courcelles, M. Desjardins, and P. Thibault. 2009. The Phagosomal Proteome in Interferon-γ-Activated Macrophages. Immunity. 30:143–154. doi:10.1016/j.immuni.2008.11.006. Ugur, B., W. Hancock-Cerutti, M. Leonzino, and P. De Camilli. 2020. Role of VPS13, a protein with similarity to ATG2, in physiology and disease. Current Opinion in Genetics and Development. 65:61–68. doi:10.1016/j.gde.2020.05.027. Valverde, D.P., S. Yu, V. Boggavarapu, N. Kumar, J.A. Lees, T. Walz, K.M. Reinisch, and T.J. Melia. 2019. ATG2 transports lipids to promote autophagosome biogenesis. The Journal of Cell Biology. 218:1787–1798. doi:10.1083/jcb.201811139. Vance, J.E., E.J. Aasman, and R. Szarka. 1991. Brefeldin A does not inhibit the movement of phosphatidylethanolamine from its sites for synthesis to the cell surface. J Biol Chem. 266:8241–8247. Varshney, P., V. Yadav, and N. Saini. 2016. Lipid rafts in immune signalling: current progress and future perspective. Immunology. 149:13–24. doi:10.1111/imm.12617. Velayos-Baeza, A., C. Levecque, C. Dobson-Stone, and A.P. Monaco. 2008. The Function of Chorein. In Neuroacanthocytosis Syndromes II. R.H. Walker, S. Saiki, and A. Danek, editors. Springer Publishing. 87–106. Velayos-Baeza, A., A. Vettori, R.R. Copley, C. Dobson-Stone, and A.P. Monaco. 2004. Analysis of the human VPS13 gene family. Genomics. 84:536–549. doi:10.1016/j.ygeno.2004.04.012. Velikkakath, A.K.G., T. Nishimura, E. Oita, N. Ishihara, and N. Mizushima. 2012. Mammalian Atg2 proteins are essential for autophagosome formation and important for regulation of size and distribution of lipid droplets. Molecular Biology of the Cell. 23:896–909. doi:10.1091/mbc.e11-09-0785. Vonk, J.J., W.M. Yeshaw, F. Pinto, A.I.E. Faber, L.L. Lahaye, B. Kanon, M. van der Zwaag, A. Velayos-Baeza, R. Freire, S.C. van IJzendoorn, N.A. Grzeschik, and O.C.M. Sibon. 2017. Drosophila Vps13 Is Required for Protein Homeostasis in the Brain. PLOS ONE. 12:e0170106. doi:10.1371/journal.pone.0170106. Wang, C., B. Wang, T. Pandey, Y. Long, J. Zhang, F. Oh, J. Sima, R. Guo, Y. Liu, C. Zhang, S. Mukherjee, M. Bassik, W. Lin, H. Deng, G. Vale, J.G. McDonald, K. Shen, and D.K. Ma. 2022. A conserved megaprotein-based molecular bridge critical for lipid trafficking and cold resilience. Nat Commun. 13:6805. doi:10.1038/s41467-022-34450-y. Wang, D.-Y., R. Fulthorpe, S.N. Liss, and E.A. Edwards. 2004. Identification of Estrogen-Responsive Genes by Complementary Deoxyribonucleic Acid Microarray and Characterization of a Novel Early Estrogen-Induced Gene: EEIG1. Molecular Endocrinology. 18:402–411. doi:10.1210/me.2003-0202. 182 Wang, J., N. Fang, J. Xiong, Y. Du, Y. Cao, and W.-K. Ji. 2021. An ESCRT-dependent step in fatty acid transfer from lipid droplets to mitochondria through VPS13D−TSG101 interactions. Nat Commun. 12:1252. doi:10.1038/s41467-021-21525-5. Wang, M.L., M. Motamed, R.E. Infante, L. Abi-Mosleh, H.J. Kwon, M.S. Brown, and J.L. Goldstein. 2010. Identification of surface residues on Niemann-Pick C2 essential for hydrophobic handoff of cholesterol to NPC1 in lysosomes. Cell Metabolism. 12:166–173. doi:10.1016/j.cmet.2010.05.016. Watanabe, Y., T. Kobayashi, H. Yamamoto, H. Hoshida, R. Akada, F. Inagaki, Y. Ohsumi, and N.N. Noda. 2012. Structure-based analyses reveal distinct binding sites for Atg2 and phosphoinositides in Atg18. Journal of Biological Chemistry. 287:31681–31690. doi:10.1074/jbc.M112.397570. Wen, P.J., S.L. Osborne, I.C. Morrow, R.G. Parton, J. Domin, and F.A. Meunier. 2008. Ca 2+ -regulated Pool of Phosphatidylinositol-3-phosphate Produced by Phosphatidylinositol 3-Kinase C2α on Neurosecretory Vesicles. MBoC. 19:5593–5603. doi:10.1091/mbc.e08-06-0595. Williamson, P. 2016. Phospholipid Scramblases. Lipid Insights. 8:41–44. doi:10.4137/LPI.S31785. Wong, L.H., A.T. Gatta, and T.P. Levine. 2019. Lipid transfer proteins: the lipid commute via shuttles, bridges and tubes. Nature Reviews Molecular Cell Biology. 20:85–101. doi:10.1038/s41580-018-0071-5. Wong, L.H., and T.P. Levine. 2016. Lipid transfer proteins do their thing anchored at membrane contact sites… but what is their thing? Biochem Soc Trans. 44:517–527. doi:10.1042/BST20150275. Wong, L.H., and T.P. Levine. 2017. Tubular lipid binding proteins (TULIPs) growing everywhere. Biochim Biophys Acta. 1864:1439–1449. doi:10.1016/j.bbamcr.2017.05.019. Wu, H., P. Carvalho, and G.K. Voeltz. 2018. Here, there, and everywhere: The importance of ER membrane contact sites. Science. 361:eaan5835. doi:10.1126/science.aan5835. Xu, P., R.D. Baldridge, R.J. Chi, C.G. Burd, and T.R. Graham. 2013. Phosphatidylserine flipping enhances membrane curvature and negative charge required for vesicular transport. J Cell Biol. 202:875–886. doi:10.1083/jcb.201305094. Yang, J., R. Yan, A. Roy, D. Xu, J. Poisson, and Y. Zhang. 2015. The I-TASSER Suite: protein structure and function prediction. Nature Methods. 12:7–8. doi:10.1038/nmeth.3213. Yang, R.-Y., H. Xue, L. Yu, A. Velayos-Baeza, A.P. Monaco, and F.-T. Liu. 2016. Identification of VPS13C as a galectin-12-binding protein that regulates galectin-12 protein stability and adipogenesis. PLoS ONE. 11. doi:10.1371/journal.pone.0153534. 183 Yang, Y., M. Lee, and G.D. Fairn. 2018. Phospholipid subcellular localization and dynamics. Journal of Biological Chemistry. 293:6230–6240. doi:10.1074/jbc.R117.000582. Yang, Y., K. Sun, W. Liu, X. Li, W. Tian, P. Shuai, and X. Zhu. 2021. The phosphatidylserine flippase β-subunit Tmem30a is essential for normal insulin maturation and secretion. Molecular Therapy. 29:2854–2872. doi:10.1016/j.ymthe.2021.04.026. Yeshaw, W.M., M. van der Zwaag, F. Pinto, L.L. Lahaye, A.I. Faber, R. Gómez-Sánchez, A.M. Dolga, C. Poland, A.P. Monaco, S.C. van IJzendoorn, N.A. Grzeschik, A. Velayos-Baeza, and O.C. Sibon. 2019. Human VPS13A is associated with multiple organelles and influences mitochondrial morphology and lipid droplet motility. eLife. 8. doi:10.7554/eLife.43561. Yochem, J., L.R. Bell, and R.K. Herman. 2004. The Identities of sym-2, sym-3 and sym-4, Three Genes That Are Synthetically Lethal With mec-8 in Caenorhabditis elegansSequence data from this article have been deposited with the EMBL/GenBank Data Libraries under accession nos. AY220985, AY221634, AY223545, and AY372076. Genetics. 168:1293–1306. doi:10.1534/genetics.104.029827. Yu, T.W., M.H. Chahrour, M.E. Coulter, S. Jiralerspong, K. Okamura-Ikeda, B. Ataman, K. Schmitz-Abe, D.A. Harmin, M. Adli, A.N. Malik, A.M. D’Gama, E.T. Lim, S.J. Sanders, G.H. Mochida, J.N. Partlow, C.M. Sunu, J.M. Felie, J. Rodriguez, R.H. Nasir, J. Ware, R.M. Joseph, R.S. Hill, B.Y. Kwan, M. Al-Saffar, N.M. Mukaddes, A. Hashmi, S. Balkhy, G.G. Gascon, F.M. Hisama, E. LeClair, A. Poduri, O. Oner, S. Al-Saad, S.A. Al-Awadi, L. Bastaki, T. Ben-Omran, A.S. Teebi, L. Al-Gazali, V. Eapen, C.R. Stevens, L. Rappaport, S.B. Gabriel, K. Markianos, M.W. State, M.E. Greenberg, H. Taniguchi, N.E. Braverman, E.M. Morrow, and C.A. Walsh. 2013. Using whole-exome sequencing to identify inherited causes of autism. Neuron. 77:259–73. doi:10.1016/j.neuron.2012.11.002. Zaman, M.F., A. Nenadic, A. Radojičić, A. Rosado, and C.T. Beh. 2020. Sticking With It: ER-PM Membrane Contact Sites as a Coordinating Nexus for Regulating Lipids and Proteins at the Cell Cortex. Frontiers in Cell and Developmental Biology. 8. Zdobnov, E.M., F. Tegenfeldt, D. Kuznetsov, R.M. Waterhouse, F.A. Simão, P. Ioannidis, M. Seppey, A. Loetscher, and E.V. Kriventseva. 2017. OrthoDB v9.1: cataloging evolutionary and functional annotations for animal, fungal, plant, archaeal, bacterial and viral orthologs. Nucleic Acids Res. 45:D744–D749. doi:10.1093/nar/gkw1119. Zecchin, A., P.C. Stapor, J. Goveia, and P. Carmeliet. 2015. Metabolic pathway compartmentalization: an underappreciated opportunity? Current Opinion in Biotechnology. 34:73–81. doi:10.1016/j.copbio.2014.11.022. Zhang, D., and L. Aravind. 2010. Identification of novel families and classification of the C2 domain superfamily elucidate the origin and evolution of membrane targeting activities in eukaryotes. Gene. 469:18–30. doi:10.1016/j.gene.2010.08.006. 184 Zhang, Z., E. Hui, E.R. Chapman, and M.B. Jackson. 2009. Phosphatidylserine Regulation of Ca 2+ -triggered Exocytosis and Fusion Pores in PC12 Cells. MBoC. 20:5086–5095. doi:10.1091/mbc.e09-08-0691. Zhong, Z., V. Pannu, M. Rosenow, A. Stark, and D. Spetzler. 2018. KIAA0100 Modulates Cancer Cell Aggression Behavior of MDA-MB-231 through Microtubule and Heat Shock Proteins. Cancers. 10:180. doi:10.3390/cancers10060180.
Thesis/Dissertation
eng
Medical Genetics
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Conibear, Elizabeth
Membrane contact site targeting and functions of Vps13 and related bridging lipid transporters
Text