Structural and Post-Transcriptional Regulation of Autophagy In

Structural and

Post-Transcriptional Regulation of Autophagy

Damián Gatica Mizala

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy (Molecular, Cellular and Developmental Biology) in the University of Michigan 2019

Doctoral Committee:

Professor Daniel J. Klionsky, Chair Professor Jayakrishnan Nandakumar Professor Laura Olsen Professor Lois Weisman

Damián Gatica Mizala

[email protected]

ORCID iD: 0000-0001-5052-3131

Dedication

To my parents Alejandra and Jaime

ii Acknowledgements

First, I would like to thank my advisor Dr. Daniel J. Klionsky not only for his amazing mentoring role in my doctoral work throughout these years, but also, and probably unbeknownst to him, for his significant impact in my life. More than 5 years ago when I was looking for a Ph.D. program I emailed several faculty members in different colleges hoping I could learn more about their work and possibly join their lab in the future. One of the few that replied was Dr. Klionsky, this very short moment of kindness that he had with someone from a third world country all across the world that he had never met before was one of the reasons I decided to apply to Michigan so I could join his lab. Looking back, I can’t but feel extremely satisfied with this decision. As a Ph.D. mentor, advisor, role model, scientist, guide and support, Dr. Klionsky has fulfilled and surpassed all my expectations many times over and for that I will always be grateful.

I am also very grateful for my thesis committee who guided me continuously since 2014.

In particular, I would like to thank Dr. Lois Weisman for her challenging questions and constructive criticism, Dr. Laura Olsen for always being on top of the next steps to follow and Dr.

J.K Nandakumar for bringing and sharing his outside expertise to the discussion.

I’m a true believer that in order to improve you need to learn from the best and in this regard, I am extremely fortunate to have joined the Klionsky lab where I was able to work with so many amazing scientist and caring people. It’s hard to think of a laboratory as a place where you

iii belong but somehow the Klionsky lab accomplishes this from the moment you enter. I sincerely thank all past and current lab members for their time, patience and encouragement that make up the fabric of the lab. I would like to thank Dr. Amélie Bernard for giving me her time and understanding both inside and outside the lab when I was just beginning my research. I would also like to thank Dr. Meiyan Jin; I will continue to miss and cherish the time we spent at the lab together. I thank Dr. Xu Liu for his patience and kindness; he was always there for me when I needed help and continuously supported me with my experiments. I would like to thank my ultra- friend Dr. Yuchen Feng for her optimism, laughter and probably being the main motivator for the lab to perform any social activity outside the building. I thank Dr. Aileen Ariosa for being such a modest and kindhearted person, always willing to provide help when needed. My most sincere gratitude to Dr. Hana Popelka for her immense scientific vision, creativity and imagination. I thank ancient Dr. Zhiyuan Yao for welcoming me to the lab with open arms. I thank Ying Yang, for her contagious joy and for being such a great pupil. I would also like to thank Vikramjit Lahiri for his sense of humor and Clarence Pascual for his enormous heart. I would also like to thank other members Vishhvaan Gopalakrishnan, Dr. Ke Wang, Dr. Katie Parzych, Daniel Orban, Dr.

Elizabeth Delorme-Axford, Zhangyuan Yin, Ziheng Xu, Wayne Hawkins, Yuchen Lei, Padma

Metur and Zhihai Zhang for generating such a positive working environment. Last but certainly not least I would like to specially thank Xin Wen, who I feel as my very own little sister and will probably only read this when she asks me for my dissertation next year. Xin, I thank you for your kindness, your smile, your resilience and for being a true friend. As I write this, I sadly realize that not long from now we will no longer be labmates. However, I want you to remember that, even if

iv we don’t see each other every day or we are on different parts of the world, no matter what happens,

I will always have your back. To all the lab, I have learned from each and every one of you and that is an honor that I will never be able to repay.

I am very thankful for the time spent with the members of my cohort which includes Jiyuan,

Nebibe, Lu, Wenjia, Yi and specially Po-Ju who has remained a good friend throughout these years.

I would also like to show my appreciation to all my friends back in Chile who, although from far away, have kept encouraging me throughout these years. In specific, I would like to thank my brothers Agustín, Alberto, Alejandro and Tomás, who I have remembered and missed every single day since I left Chile. I would also like to give a big hug to Carmen, Fernanda, Sofía,

Valentina and specially Paula for their efforts in keeping up with someone you barely see once a year and yet you keep calling a friend like it was only yesterday we saw each other for the last time. Special thanks also go out to Sebastián who time and time again has proven to be an endless source of wisdom, laughter and friendship.

My deepest gratitude goes to who I have no doubt will become my future father and mother in law, Guillermo and Eleana, who have consistently showed me their love and made me part of their family way before that was even a possibility. On the same note, I would like to give special thanks to Javiera, who even throughout all these years, all the questions, all the distance and all our worries is still standing by my side. Believing in me and the promise that someday we will accomplish all our dreams. I love you, I don’t know what I would do without you.

v Finally, I would like to thank my grandfather Moises, my sister Camila, my mamadre Olga and to whom I dedicate this dissertation, my parents Alejandra and Jaime. Even though I’m not a parent, as I have grown older I have learned to recognize the difficulties that come from being one.

I have looked through my memories for reasons that have led me down this path in life, finding small gifts related to science, encouragement in learning biology and chemistry; and huge smiles when I decided to study biochemistry and later apply for a Ph.D. Anyone who read this might think that I was led down this path against my own will, that I was being guided towards this because it’s what my parents wanted of me. To those people I say, you know nothing about parenting, for what I have found walking on this road, it’s only joy. My parents did everything they did not because they wanted anything, but because they listened. They heard me talking about space, science, cells and atoms, even when I was too young to even know what those things truly were, and they thought, this is what makes him happy. They helped me walk this path because they knew it would make me whole. As I grow older I can clearly see that now, and I only hope that if I ever have kids I could be half as good as they were to me. Thank you.

vi Preface

This thesis summarizes the research projects I have participated in while working in Dr.

Daniel J. Klionsky’s laboratory since joining in 2014. Through these projects we gain a better understanding of the post-transcriptional regulation of different autophagy-related (ATG) gene mRNA transcripts and new insights into the structural importance of the essential autophagy protein Atg13.

Chapter 1 contains parts of two review papers published in Circulation Research (doi: https://doi.org/10.1161/CIRCRESAHA.114.303788) and Nature Cell Biology (doi: https://doi.org/10.1038/s41556-018-0037-z) with some modifications.

Chapter 2 characterizes a new post-transcriptional pathway regulating autophagy involving the Pat1-Lsm complex and how binding of this complex facilitates ATG mRNA stabilization during nitrogen starvation-induced autophagy. This research was published in the journal

Molecular Cell (doi: https://doi.org/10.1016/j.molcel.2018.11.002). Guowu Hu and Nannan

Zhang from Peter R. Williamson’s laboratory in the National Institutes of Health provided mammalian data. Xu Liu helped optimize RNA-protein Bimolecular Fluorescence

Complementation and RNA immunoprecipitation assays. All other data were generated by me.

Chapter 3 characterizes an undiscovered binding mechanism between the essential autophagy protein Atg13’s intrinsically disordered carboxy terminus region and lipid membranes.

This manuscript is submitted and under revision. Hana Popelka and Clarence Pascual helped with experiment design, performed strain construction and affinity isolation. Alejandro Damasio from

Michael J. Ragusa’s laboratory in Dartmouth College performed 1D 1H NMR spectroscopy,

vii circular dichroism, liposome sedimentation assays and isothermal titration calorimetry. All other data were generated by me.

Chapter 4 summarizes the major implications of my thesis research and future directions.

viii Table of Contents

Dedication ...... ii

Acknowledgements ...... iii

Preface ...... vii

List of Tables ...... xi

List of Figures ...... xii

Abstract ...... xv

Chapter 1 Introduction ...... 1

1.1 Overview of Autophagy ...... 1 1.2 Membrane nucleation and source ...... 5 1.3 Phagophore expansion ...... 9 1.4 Lysosome/Vacuole fusion and recycling of degraded cargo ...... 11 1.5 Selective autophagy overview: Cytoplasm-to-vacuole targeting pathway ...... 11

Chapter 2 The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen

Starvation-Induced Autophagy ...... 39

2.1 Abstract ...... 39 2.2 Introduction ...... 40 2.3 Results ...... 41 2.4 Discussion ...... 54 2.5 Materials and Methods ...... 57 2.6 Acknowledgments ...... 64

ix Chapter 3 The carboxy terminus of yeast Atg13 binds phospholipid membrane via motifs that overlap with the Vac8-interacting domain ...... 86

3.1 Abstract ...... 86 3.2 Introduction ...... 87 3.3 Results ...... 89 3.4 Discussion ...... 100 3.5 Materials and Methods ...... 102 3.6 Acknowledgements ...... 106

Chapter 4 Summary ...... 128

4.1 The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen Starvation-Induced Autophagy ...... 129 4.2 The carboxy-terminus of yeast Atg13 binds phospholipid membrane via motifs that overlap with the Vac8-interacting domain ...... 131 4.3 Discussion and perspectives ...... 133

Bibliography ...... 135

x List of Tables

Table 1.1 Selective autophagy ligands, receptors and scaffolds in yeast and mammals

...... 28

Table 2.1 S. cerevisiae strains used in this study...... 65

Table 2.2 Primers used in this study ...... 67

Table 3.1 S. cerevisiae strains used in this study...... 107

Table 3.2 Dissociation constants for the Atg13[571-700] wild-type (WT) and mutant proteins titrated into Vac8[10-515] as determined by ITC...... 109

xi List of Figures

Figure 1.1 The 3 main types of autophagy in yeast and mammals ...... 29

Figure 1.2 Autophagy induction...... 30

Figure 1.3 Autophagy regulation ...... 31

Figure 1.4 Class III phosphatidylinositol 3-kinase (PtdIns3K) complexes...... 32

Figure 1.5 Autophagosomes have a diverse range of potential membrane sources . 33

Figure 1.6 Two ubiquitin-like conjugation systems ...... 34

Figure 1.7 The Cvt pathway and aggrephagy ...... 35

Figure 1.8 Pexophagy ...... 36

Figure 1.9 Mitophagy, Reticulophagy and Nucleophagy ...... 37

Figure 1.10 Lysophagy and Xenophagy ...... 38

Figure 2.1 PAT1 Deletion Lowers Autophagy Activity after Nitrogen starvation by

Decreasing ATG mRNA and Protein Levels ...... 69

Figure 2.2 LSM1 deletion lowers autophagy activity after nitrogen starvation by decreasing ATG mRNA and protein levels ...... 70

Figure 2.3 SKI3 deletion prevents the pat1∆-mediated decrease in autophagy ...... 71

Figure 2.4 SKI3 deletion prevents the lsm1∆-mediated decrease in autophagy ...... 73

xii Figure 2.5 Pat1 is dephosphorylated under nitrogen-starvation conditions ...... 74

Figure 2.6 Pat1 dephosphorylation on S456 and S457 regulates autophagy by modulating ATG mRNA and protein levels during nitrogen starvation ...... 75

Figure 2.7 Pat1 binds specific ATG mRNAs ...... 76

Figure 2.8 Protein-RNA BiFC of ATG2-MS2, ATG17-MS2 and ATG18-MS2...... 78

Figure 2.9 SKI3 deletion prevents the Pat1S456E- and Pat1S457E-mediated decrease in autophagy ...... 79

Figure 2.10 Switching the ATG1 mRNA 3' UTR prevents the PAT1 deletion-mediated decrease in ATG1 mRNA and protein levels ...... 81

Figure 2.11 PATL1 levels and phosphorylation state regulate ATG2 and ATG9A mRNA levels and autophagy activity...... 83

Figure 2.12 Pat1 stabilizes a subset of ATG mRNAs during nitrogen starvation- induced autophagy by preventing 3'-5' mRNA degradation ...... 84

Figure 3.1 1D 1H NMR spectrum of Atg13[571-700] ...... 110

Figure 3.2 Circular dichroism ...... 110

Figure 3.3 Atg13[571-700] binds negatively charged membranes ...... 111

Figure 3.4 Autophagy activity of Atg13 mutants ...... 113

Figure 3.5 Atg13 utilizes two distinct regions for liposome binding ...... 115

Figure 3.6 Schematic illustration of Atg13 mutants and summary of liposome binding assays ...... 117

xiii

Figure 3.7 Atg13 IDR mutants reduce autophagy activity ...... 118

Figure 3.8 Atg13[571-700] interacts with Vac8 ...... 119

Figure 3.9 Interaction between Atg13 and Vac8 ...... 121

Figure 3.10 Vac8 inhibits Atg13[571-700] liposome binding ...... 123

Figure 3.11 Autophagy activity and cellular localization of the truncated Atg13 [1-

570] ...... 125

Figure 3.12 Model of proposed mechanism for a reversible switch in binding of the

Atg13 C terminus from Vac8 to phospholipid membranes ...... 127

xiv Abstract

Autophagy is a highly conserved pathway in eukaryotes, involving cellular recycling of multiple cytoplasmic components during standard physiological conditions and in response to different types of stress, such as nutrient starvation. Macroautophagy/autophagy can be either non-selective or selective and involves the sequestration of cytoplasm within double-membrane vesicles termed autophagosomes. Upon maturation, autophagosomes fuse with the vacuole (in yeast and plants) or lysosomes (in metazoans), leading to degradation of their cargo by resident hydrolases. Cargo degradation produces basic molecules such as amino acids, which are subsequently recycled back into the cytoplasm for reuse. Whereas non-selective autophagy, a cellular response to nutrient deprivation, typically involves random uptake of cytoplasm by the membrane precursors to autophagosomes, termed phagophores, selective autophagy is responsible for specifically removing certain components such as protein aggregates and damaged or superfluous organelles.

Autophagy dysregulation has been involved in the development of a diverse range of diseases such as cancer and different neurodegenerative disorders. Several autophagy-related (ATG) genes and their corresponding proteins have been characterized based on their function(s) in regulating the various stages of autophagy. Upon nutrient starvation ATG gene transcription is dramatically upregulated to increase Atg protein synthesis and autophagy activity. However, we are still understanding the different types of regulation involved in autophagy signaling both at the post- transcriptional and structural level; and how mutations in these ATG genes can lead to different types of disease.

xv Pat1 is an mRNA binding protein that interacts with the heptameric ring-shaped complex formed by Lsm1 to Lsm7, and together forms the Pat1-Lsm complex. Although initially characterized as a decapping-enhancing factor, we showed that during nitrogen starvation the Pat1-

Lsm complex can bind and stabilize a specific subset of ATG mRNA by preventing their 3’ to 5’ degradation by the exosome exonuclease complex. We provide evidence that this process is regulated through Pat1 dephosphorylation, is necessary for the efficient expression of specific Atg proteins and is required for normal autophagy induction during nitrogen starvation.

The essential autophagy protein Atg13 is a key member of the autophagy initiation complex. Atg13 is thought to work as a hub for assembly between the initiation complex and participating membranes. The Atg13 C terminus is predicted to be an intrinsically disordered region (IDR) harboring a binding site for the vacuolar membrane protein Vac8. We discovered that Atg13 C terminus IDR directly binds to lipid membranes via electrostatic interactions between positively charged residues in Atg13 and negatively charged phospholipids. We identified 2 sets of residues in the Atg13 IDR that affect its phospholipid binding properties; and these residues overlap with the Vac8-binding domain of Atg13. Our data indicate that Atg13 binding to phospholipids and Vac8 is mutually exclusive, and both are required for efficient autophagy.

The research presented in this dissertation gives an overview of the different mechanisms by which autophagy can be regulated both at the mRNA and structural protein level.

xvi Chapter 1 Introduction1

1.1 Overview of Autophagy

1.1.1 Definition

Autophagy is a highly conserved catabolic pathway in eukaryotes, involving the cellular recycling of multiple cytoplasmic components during standard physiological conditions and in response to different types of stress, such as nutrient starvation1. Autophagy can be divided into 3 main types: macroautophagy, microautophagy, and chaperone-mediated autophagy. Whereas microautophagy and macroautophagy can be selective or nonselective processes found in yeast and higher eukaryotes (Figure 1.1A), chaperone-mediated autophagy is a selective process that has only been described in mammalian cells2. During chaperone-mediated autophagy, specific protein substrates containing the amino acid sequence KFERQ are recognized by chaperones, unfolded and translocated into the lysosome through the lysosomal membrane protein LAMP2A (Figure 1.1B)3.

During microautophagy, uptake also occurs directly at the limiting membrane of the lysosome (in metazoans) lysosome or vacuole (in yeasts and plants). In this case, however, the process operates by directly sequestering the substrates through invagination of the or vacuole/lysosome membrane4. Of the 3 types mentioned, macroautophagy (hereafter autophagy) is clearly the most well-studied process. Autophagy involves the sequestration of cytoplasm within double-membrane

1 This chapter is reprinted partly from Damian Gatica, Mario Chiong, Sergio Lavandero, Daniel J. Klionsky (2015) Circulation Research (doi: https://doi.org/10.1161/CIRCRESAHA.114.303788), and partly from Damian Gatica, Vikramjit Lahiri, Daniel J Klionsky (2018) Nature Cell Biology (doi: https://doi.org/10.1038/s41556-018-0037-z), with minor modifications.

1 vesicles termed autophagosomes. Upon maturation, autophagosomes fuse with the vacuole or lysosomes, leading to degradation of their cargo by resident hydrolases1, 5. Cargo degradation produces basic molecules such as amino acids, which are subsequently recycled back into the cytoplasm for reuse1, 6. Whereas non-selective autophagy, a cellular response to nutrient deprivation, typically involves random uptake of cytoplasm into the precursor of autophagosomes, termed phagophores, selective autophagy is responsible for specifically removing certain components such as protein aggregates and damaged or superfluous organelles7. Different studies have reported the selective autophagic degradation of several organelles, including mitochondria8, peroxisomes9, lysosomes10, endoplasmic reticulum (ER) and the nucleus11, under various conditions. Furthermore, autophagy selectively degrades aggregation-prone misfolded proteins and protein microaggregates implicated in the pathology of various neurodegenerative diseases12.

Multiple autophagy-related (ATG) genes and proteins have been described as being involved in the different stages of autophagy, comprising what is now known as the core autophagy machinery that is required for autophagosome formation, and additional proteins that act in making the process selective, or in stages other than autophagosome biogenesis. Accordingly, autophagy can be dissected into different steps based on the proteins involved, including induction, nucleation of the phagophore, membrane expansion and maturation of the autophagosome, fusion with the lysosome/vacuole, and recycling of the degraded cargo.

1.1.2 Autophagy induction in yeast

During growth in nutrient-rich conditions autophagy activity is kept to a minimum by different nutrient sensing pathways including those regulated by the target of rapamycin (TOR) kinase and cAMP-dependent protein kinase A. The ability of TOR to sense nutrient levels, in particular amino acids, makes it a critical negative regulator of autophagy. The rapamycin sensitive TOR complex

2 1 (TORC1) inhibits autophagy in part by preventing the activation of the Atg1 kinase complex13.

In yeast, the Atg1 kinase complex is formed by the Ser/Thr kinase Atg1, the regulatory subunit protein Atg13, and the Atg17–Atg31–Atg29 complex, which is thought to function as a scaffold13,

14. Although specific substrates of Atg1 are still being discovered, the Atg1 kinase complex plays an essential role in autophagy induction by recruiting other Atg proteins to what is known as the phagophore assembly site (PAS), a perivacuolar location found in yeast that is proposed to be the organizing center for phagophore formation15-18. Upon nutrient starvation or rapamycin treatment

TORC1 activity is inhibited and Atg13 is rapidly but partially dephosphorylated leading to the activation of Atg119. Autophosphorylation of Atg1 within its activation loop is also important for activating its kinase activity and inducing autophagy (Figure 1.2A)20. Atg13 dephosphorylation was linked to an increased interaction with Atg1 leading to a model in which starvation increased affinity between the 2 proteins; however, recent data as well as interaction studies in other model organisms support the idea that Atg1 and Atg13 interact independently of nutrient conditions21.

In addition to Atg13, TORC1 may also inhibit autophagy by directly phosphorylating Atg122.

Protein kinase A is also a negative regulator of autophagy. Protein kinase A suppresses autophagy by phosphorylating both Atg1 and Atg1323-25. Although the 2 pathways target similar proteins,

TOR and protein kinase A seem to work largely independent of one another by targeting different phosphorylation sites25. Other nutrient-sensing kinases involved in autophagy induction include

Snf1 and Gcn2. Snf1 corresponds to the yeast homolog of 5′-AMP–activated protein kinase

(AMPK), which will be discussed below. Gcn2 promotes autophagy during amino acid starvation by phosphorylating Sui2/eIF2α (suppressor of initiator codon 2). Phosphorylation of Sui2 blocks general protein synthesis and specifically activates the translation of the transcription factor Gcn4, which in turn induces the transcription of various ATG genes26, 27.

3 1.1.3 Autophagy induction in Mammals

In contrast to yeast, mammalian cells have multiple Atg1 homologs, and the ones most relevant to autophagy are unc-51 like autophagy activating kinase 1 (ULK1) and ULK228, 29. Thus, in mammals the Atg1 kinase complex is known as the ULK kinase complex and is formed by

ULK1/2, the mammalian homolog of Atg13 (ATG13), the functional homolog of Atg17

(RB1CC1), and the ATG13-stabilizing protein ATG101, which has no yeast counterpart. All members of the ULK kinase complex are required for autophagy induction in mammalian cells30-

32. As mentioned above, in mammalian cells the interaction between the members of the ULK kinase complex does not depend on nutrient conditions33. Although some studies indicate that

ATG13 mediates the interaction between RB1CC1 and ULK31, others have reported that all members of the complex can interact independently33. Similar to the yeast Atg1 complex, regulation of the ULK kinase complex depends on the mechanistic target of rapamycin complex

1 (MTORC1). During nutrient-rich conditions, MTORC1 interacts directly with ULK1 through the scaffold protein regulatory associated protein of MTOR, complex 1 (RPTOR) and inhibits its kinase activity by phosphorylating both ATG13 and ULK1/231, 34. Upon nutrient starvation or rapamycin treatment, MTORC1 is released from the ULK kinase complex leading to the dephosphorylation of both proteins and the activation of ULK kinase activity31, 33, 34. Once activated, ULK1 phosphorylates ATG13, RB1CC1, and itself, stabilizing its enzymatic activity and inducing the autophagic process (Figure 1.2B)31, 34, 35. Another protein capable of sensing energy levels that is involved in autophagy regulation is AMPK. Through the upstream kinase serine/threonine kinase 11 (STK11/LKB1), AMPK is able to sense decreases in the cellular

ATP/AMP ratio leading to its activation and autophagy induction36. During glucose deprivation

AMPK phosphorylates and activates the tuberous sclerosis complex, tuberous sclerosis (TSC) 1-

4 TSC2, which in turn inactivates the GTPase-activating protein Ras homolog enriched in brain

(RHEB), leading to MTORC1 inhibition and the release of the ULK kinase complex (Figure 1.3)37,

38. Once MTORC1 leaves the ULK complex, AMPK directly phosphorylates ULK1, stimulating its catalytic activity and inducing autophagy36. Interestingly, the ULK kinase complex also phosphorylates and inactivates AMPK, through a mechanism that has been described as an inhibitory feedback loop39. Although functioning in part in a hormone-sensing pathway,

AKT/PKB can also regulate autophagy by controlling MTORC1 activation. On ligand binding, dimerization, autophosphorylation, and activation of insulin receptor or insulin-like growth factor

1 receptor, the class I phosphoinositide 3-kinase is recruited to the plasma membrane and activated40. Phosphoinositide 3-kinase catalyzes the phosphorylation of phosphatidylinositol (4,5) bisphosphate generating the lipid second messenger phosphatidylinositol (3,4,5) trisphosphate

(PIP3), which in turn recruits AKT to the plasma membrane where it is activated via phosphorylation by PDPK1 and MTORC240, 41. AKT-dependent phosphorylation of TSC2 prevents RHEB inhibition, leading to MTORC1 activation and autophagy inhibition42, 43. As a consequence, the tumor suppressor and lipid phosphatase and tensin homolog (PTEN) can induce autophagy by dephosphorylating PIP3 and downregulating the AKT phosphoinositide 3-kinase pathway (Figure 1.3)44.

1.2 Membrane nucleation and source

Once autophagy is induced, assembly of the phagophore is initiated by membrane nucleation. As mentioned above, in yeast the PAS corresponds to the location at which several Atg proteins are recruited to assemble the phagophore. In contrast, mammalian cells lack a single defined PAS, and autophagosome formation seems to be initiated at different locations inside the cell. In both yeast and mammals, the class III phosphatidylinositol 3-kinase (PtdIns3K) catalyzes the nucleation of

5 the phagophore by generating phosphatidylinositol 3-phosphate (PtdIns3P) and inducing the recruitment of PtdIns3P-binding proteins43. In yeast the PtdIns3K is formed by the regulatory subunit vacuolar protein sorting 15 (Vps15), the catalytic subunit Vps34, Vps30/Atg6, Atg14 and

Atg38, all of which are essential for autophagy45-47. Similarly, the core mammalian PtdIns3K is composed of the Vps15 homolog phosphoinositide-3-kinase, regulatory subunit 4 (PIK3R4), the

Vps34 homolog phosphatidylinositol 3-kinase, catalytic subunit type 3 (PIK3C3), and the

Vps30/Atg6 homolog beclin-1, autophagy related (BECN1)43, 48. Although these 3 proteins constitute the core machinery of the mammalian PtdIns3K, distinct interactions with specific proteins lead to the formation of ≥3 different PtdIns3K complexes that play different roles in autophagy49-51. One of these complexes is formed by the interaction of the PtdIns3K core complex with the mammalian Atg14 homolog (ATG14) and autophagy/beclin-1 regulator 1 (AMBRA1)52,

53. The ATG14-containing PtdIns3K complex is thought to positively regulate autophagy by promoting translocation of the complex to the phagophore and inducing the generation of

PtdIns3P50, 53, 54. The other 2 PtdIns3K complexes contain the BECN1-interacting protein UV radiation resistance associated (UVRAG) as a common component. Whereas the PtdIns3K complex formed by UVRAG and SH3-domain GRB2-like endophilin B1 (SH3GLB1/Bif-1) promotes autophagosome formation54, 55, the complex formed by UVRAG and

KIAA0226/RUBICON downregulates autophagy by impairing autophagosome maturation

(Figure 1.4)50, 51. Other BECN1-interacting proteins include the antiapoptotic protein B-cell

CLL/lymphoma 2 (BCL2), which inhibits the PtdIns3K complex by sequestering BECN1 under nutrient-rich conditions56. Besides alterations in protein interactions, BECN1 post-translational modifications also regulate PtdIns3K activity. For example, BECN1 phosphorylation by DAPK promotes dissociation of BCL2 and autophagy induction57. ULK1-dependent phosphorylation of

6 BECN1 activates the ATG14- and UVRAG-containing PtdIns3K complexes inducing autophagy during amino acid starvation58. Activation of both these PtdIns3K complexes by ULK1-mediated

BECN1 phosphorylation would argue for the importance of this post-translational modification for autophagosome induction and later maturation and provides a link between the ULK kinase initiation complex and the membrane nucleation complex. Most recently, AMPK was described as regulating the activity of different PtdIns3K complexes by phosphorylating BECN1 and

PIK3C359. Although membrane nucleation has been established as a key step in the autophagic process, the origin of the membrane that gives rise to the phagophore, and subsequently the autophagosome, remains an open question. Different studies have described the ER, mitochondria, plasma membrane, and trans-Golgi network as possible membrane donors (Figure 1.5)60-62.

Evidence supporting the ER as a possible membrane source include 3-dimensional tomography studies showing a connection between the phagophore and ER, as well as ATG14-containing

PtdIns3K complex localization to the ER to initiate autophagosome formation54, 63. Generation of

PtdIns3P at the ER triggers the recruitment of the PtdIns3P-binding protein ZFYVE1/DFCP1 (zinc finger, FYVE domain containing 1) and one of the mammalian homologs of Atg18, WD repeat domain, phosphoinositide interacting 2 (WIPI2). Both of these proteins have been linked to autophagosome formation from a PtdIns3P-enriched ER-associated structure termed the omegasome for its Ω-like shape64, 65. Omegasomes have been described as platforms for autophagome formation, which seem to depend on PtdIns3P, because ATG14 depletion leads to omegasome disappearance54. Although the role of ZFYVE1 in autophagy is not well defined,

WIPI2 silencing results in accumulation of omegasome structures and failure to mature into autophagosomes, suggesting that WIPI2 is involved in the transition between omegasomes and autophagosomes65. Other Atg proteins that have been associated with omegasomes include the

7 ULK kinase complex, which localizes transiently to omegasomes in a PtdIns3P-dependent manner64, 66. The precise mechanism by which the ER gives rise to autophagosomes via an omegasome intermediate is unknown, and several questions remain to be answered about the conditions, specific proteins involved, and the selectivity of the process (Figure 1.5)67. As mentioned above, mitochondria are another organelle that have been proposed as a membrane source for the phagophore. During starvation conditions an outer mitochondria membrane fluorescent marker colocalizes with autophagosomes; mitochondrial lipids also seem to transit to autophagosomes60. The same study showed that autophagosome formation during nutrient starvation is impaired in cells lacking the ER-mitochondria tethering protein MFN2 (mitofusin 2).

Mitochondria-associated ER membrane, which are sites where the mitochondria and ER are in close proximity to each other, have been implicated in autophagosome formation. ATG14 and other autophagy markers localize to the mitochondria-associated ER membrane during starvation conditions. In MFN2–depleted cells, which are unable to tether the ER to the mitochondria,

ATG14 localization to the mitochondria-associated ER membrane is impaired. In addition, the omegasome protein marker ZFYVE1 localizes to the mitochondria-associated ER membrane upon starvation68. This remarkable finding opens the possibility that the functions of omegasomes and mitochondria in autophagosome formation are essentially one and the same, unified by the association between the 2. Other studies on the transmembrane protein Atg9 have advanced our understanding of the membrane source from which phagophores are assembled. Atg9 has been characterized as a self-interacting protein containing 6 putative transmembrane domains, with both its carboxyl and amino termini facing the cytosol69, 70. In yeast, Atg9 cycles from the PAS to peripheral membranes; Atg9-containing vesicles are thought to be part of the initial membranes that will generate the phagophore71, 72. The Atg9-containing membrane reservoir seems to be

8 composed of tubules and vesicle clusters formed through the ER-Golgi trafficking pathway71; however, Atg9 also cycles between perimitochondrial sites and the PAS73. Although the precise mechanisms by which Atg9 cycling is still being studied, several Atg proteins are involved in the regulation of Atg9 movement. Atg9 anterograde transport, which is defined as movement from the peripheral sites to the PAS, depends on Atg11, Atg23, and Atg2717, 74, 75, whereas retrograde transport, that is from the PAS to the peripheral sites, is directed by the Atg1-Atg13 complex,

Atg2, Atg18, and the PtdIns3K complex17. In mammalian cells, nutrient starvation induces ATG9 redistribution from the trans-Golgi network to phagophores. Both ULK1 silencing and PtdIns3K inhibition block ATG9 trafficking to phagophores, suggesting both complexes are involved in mammalian ATG9 cycling62, 76. The MAPK pathway is also implicated in mammalian ATG9 traffic, SUPT20H/FAM48A/p38IP (suppressor of Ty 20 homolog [Saccharomyces cerevisiae]) interacts with ATG9 and induces its redistribution leading to autophagy activation. Conversely, binding between SUPT20H and MAPK14/p38α inhibits ATG9 interaction with SUPT20H and autophagy77.

1.3 Phagophore expansion

Elongation and expansion of the phagophore membrane are key steps in the autophagic process. The Atg12–Atg5-Atg16 and Atg8 conjugation systems, 2 inter-related ubiquitin-like conjugation pathways, regulate this stage in both yeast and mammals. Before being covalently linked to their final substrates, both Atg12 and Atg8 go through an activation and conjugation reaction, triggered by an E1-like and an E2-like enzyme, respectively. In the Atg12–Atg5-Atg16 system, Atg12 is first activated in an ATP-dependent manner by the E1-like activating enzyme

Atg7, forming a thioester bond between the 2 proteins78, 79. After this event, Atg12 is transferred to the E2- like conjugating enzyme Atg10, generating the Atg12–Atg10 intermediate through the

9 formation of another thioester bond80. Finally, Atg12 is covalently attached to a specific lysine residue on Atg5 in a process that, unlike ubiquitination, seems to be constitutive, irreversible and does not require an E3-like ligase enzyme81, 82. Further interaction between Atg12–Atg5 and Atg16 leads to the formation of the Atg12–Atg5-Atg16 complex. Unlike Atg12, Atg16 is not covalently bound to Atg5 and is able to self-interact when bound to Atg12–Atg5 forming a large multimeric protein complex83-85. The Atg12–Atg5-Atg16 complex is essential for autophagy and localizes to the phagophore84, 86, 87. In yeast the second ubiquitin-like conjugation system catalyzes the lipidation of Atg8 by covalently linking it to phosphatidylethanolamine (PE). The first event in this process corresponds to the cleavage of the carboxyl terminus of Atg8 by the cysteine protease

Atg4, exposing a glycine residue88, 89. In the next step, Atg7, again working as an E1-like enzyme, activates Atg890. The activated protein is then conjugated to the E2-like enzyme Atg3, before finally being linked to PE through an amide bond90. Different studies have proposed that the E3- like enzyme that facilitates the Atg8–PE linkage is the Atg12–Atg5-Atg16 complex91-94. In its conjugated form Atg8 is bound to both sides of the autophagosome membrane and thus its N terminus GFP-tagged form is widely used as an autophagosome marker95, 96. However, Atg8 lipidation is a reversible process because Atg8–PE bound to the external autophagosome membrane can be cleaved by Atg4, releasing it from the autophagosome (Figure 1.6)88.

Both ubiquitin-like conjugation pathways are conserved and work similarly between mammals and yeast with the specific difference being that mammalian cells have several Atg8 homologs further divided into the MAP1LC3/LC3 (microtubule-associated protein 1 light chain 3) and GABARAP (GABA type A receptor-associated protein) subfamilies. Although all of the homologs go through a similar conjugation process, each subfamily works at different stages of

10 autophagy97-99; the LC3 subfamily is involved in expansion of the phagophore and the GABARAP subfamily participates at a later stage in autophagosome maturation100.

1.4 Lysosome/Vacuole fusion and recycling of degraded cargo

The fusion of autophagosomes with lysosomes/vacuoles results in the generation of autolysosomes in higher eukaryotes and autophagic bodies in yeast. In either case, the fusion process seems to involve similar machinery that plays a role in other transport processes that terminate at these degradative organelles101. In yeast, this machinery includes the class C Vps/HOPS complex, the

SNARE family proteins Ykt6, Vti1, Vam3, and Vam7, the small GTPase Ypt7, and the proteins

Mon1 and Ccz1101-107. Fewer details are known in mammalian cells; however, the Ypt7 homolog

RAB7 is required108. One difference between yeast and mammals is that there is a clear convergence between autophagy and the endocytic pathway in the latter; autophagosomes can fuse with endosomes to form amphisomes that subsequently fuse with the lysosome109. Once fusion occurs, the inner autophagosomal membrane and its cargo are degraded inside the lysosome/vacuole by various hydrolases. The resulting macromolecules such as amino acids that are obtained after cargo degradation are transported back into the cytoplasm for recycling. In yeast this process is regulated by protein permeases such as Atg22110.

1.5 Selective autophagy overview: Cytoplasm-to-vacuole targeting pathway

The cytoplasm-to-vacuole targeting (Cvt) pathway is a biosynthetic autophagy-related process specific to yeast, in which vacuolar enzymes are transported from the cytoplasm into the vacuole utilizing the autophagic machinery. Among the enzymes that utilize the Cvt pathway are Ape1

(aminopeptidase I), Ape4 and Ams1 (α -mannosidase)111. Ape1 is first synthesized in the cytoplasm as an inactive proenzyme (prApe1). Following oligomerization, prApe1 is selectively recognized by the non-core, autophagy-related (Atg) protein Atg19, which functions as a receptor

11 for Ams1, prApe1 and Ape4112, 113. Once prApe1–Atg19 (the Cvt complex) is formed, Atg19 binds to the scaffold protein Atg11, which in turn directs the Cvt complex to the perivacuolar location termed the PAS. Here, autophagosomes and Cvt vesicles are formed in yeasts114, 115; interaction of

Atg19 with Atg11 is facilitated by Hrr25-dependent phosphorylation of the receptor116. After reaching the PAS, Atg19 interacts with the ubiquitin-like protein Atg8114. During autophagy and the Cvt pathway, Atg8 is covalently conjugated through its C terminus to PE. Thus, Atg8–PE is present on both the inner and outer membrane of forming autophagosomes90 (Figure 1.7A). Atg8 has been implicated in phagophore expansion and autophagosome size regulation117. Atg19 binding to Atg8 therefore tethers the Cvt complex to the Atg8–PE-conjugated sequestering vesicles. Once fully matured, Cvt vesicles fuse with the vacuole and deliver prApe1, which is then processed into its active form by resident hydrolases.

Using the Cvt pathway as a model for selective autophagy, we can propose that although the core autophagy machinery directs phagophore membrane expansion and vesicle formation, cargo selectivity is achieved by a ligand receptor and a scaffold protein-roles taken by Atg19 and

Atg11, respectively, in the Cvt pathway. Atg19 has a paralog, Atg34 (also phosphorylated by

Hrr25), which functions as an Ams1-receptor during nitrogen starvation118. Other types of selective autophagy in yeast, such as mitophagy119 and pexophagy120, 121, also rely on Atg11 as a scaffold for cargo delivery to the PAS. However, a counterpart to Atg11 has yet to be discovered in mammals. Similarly, most types of selective autophagy require binding of the cargo receptor to the core autophagy machinery. In the Cvt pathway, this process is illustrated by Atg19 binding to

Atg8 through a specific WXXL motif found on the Atg19 C terminus, similar to that seen in mammalian SQSTM1 (also known as p62)122, 123. This interaction is evolutionarily conserved, as several proteins in yeasts and more complex eukaryotes contain Atg8-interacting motifs (AIMs)

12 or LC3-interacting regions (LIRs), respectively. The AIMs or LIRs provide selective binding to yeast Atg8 or one of the members of the LC3/GABARAP family of Atg8 mammalian homologs124.

Multiple examples of scaffold and receptor proteins will be showcased as different types of selective autophagy are discuss (Table 1.1).

1.5.1 Aggrephagy

The selective degradation of protein aggregates by autophagy is known as aggrephagy. Multiple aggregation-prone proteins such as amyloid-β125, HTT (huntingtin)126 and SNCA (synuclein alpha)127 are autophagy substrates. In yeast, Cue5 is a cargo receptor for the clearance of aggregation-prone polyglutamine (polyQ)-containing proteins. Cue5 possesses a ubiquitin-binding

CUE domain and an AIM, mediating the interaction between ubiquitinated cargo and Atg8128.

Overexpression of TOLLIP, a Cue5 human homolog that also has a CUE domain, leads to degradation of polyQ protein aggregates in human cell lines129 (Figure 1.7B). Ubiquitination of substrates has been demonstrated as a key mediator in the recognition and degradation of these proteins by selective autophagy130. At least three additional mammalian cargo receptors,

SQSTM190, 123, NBR1131 and OPTN132, act as ubiquitin-binding proteins that mediate the interaction between ubiquitinated proteins and the core autophagy machinery. All three receptors possess LIRs and ubiquitin-binding domains, thus working as a bridge between LC3/GABARAP family members and ubiquitinated substrates90, 123-128, 130-134.

The nucleocytoplasmic shuttling protein WDFY3 (also known as ALFY) has been proposed as a scaffold in aggrephagy135. While unable to directly interact with ubiquitinated substrates, WDFY3 binds the core autophagy protein Atg5, the cargo receptor SQSTM1136,

GABARAP subfamily members137, and PtdIns3P, which as previously mentioned is a prominent lipid in the regulation of autophagosome membrane formation138. WDFY3-depletion hinders

13 clearance of aggregated polyQ proteins. The latter observation, in conjunction with its high number of interacting partners, suggests that WDFY3 is an important scaffold protein in SQSTM1- dependent degradation of ubiquitinated aggregates by selective autophagy. Ubiquitination plays an important role not only in substrate recognition and degradation by the ubiquitin-proteasome system (UPS), but also by selective autophagy—raising a set of questions regarding the hierarchy between these two degradation pathways. It has been proposed that protein aggregates that cannot be degraded by the UPS (for example, due to size) may be cleared by autophagy139, 140. At the same time, the lysine residues used for linkage, as well as the length and nature of the ubiquitin chains, have been proposed as a mechanism to select which degradation pathway is chosen139. However, a recent publication emphasizes the role of receptor oligomerization over the type of ubiquitination in selecting a degradation pathway141. This finding agrees with data showing the importance of

Cue5 and SQSTM1 oligomerization in their association with the phagophore139, 142. Thus, both autophagy and the UPS provide dynamic alternatives to different cellular challenges.

1.5.2 Pexophagy

Pexophagy is the selective removal of peroxisomes. Pexophagy has been mostly studied as a pathway for the removal of superfluous organelles in various fungi143. Incubating these fungi in oleic acid or methanol leads to peroxisome proliferation; following a shift to a preferred carbon source such as glucose, the excess peroxisomes are rapidly degraded through pexophagy143.

Similar to other types of selective autophagy, cargo selectivity is provided by receptor proteins; in yeast, this role is taken by Pichia pastoris Atg30 (PpAtg30)120, and by Atg36 in Saccharomyces cerevisiae144. Both Atg36 and PpAtg30 tether peroxisomes targeted for degradation to nascent phagophore membranes by linking Atg8 to peroxisomal membrane proteins, with Atg36 binding

Pex3 and PpAtg30 binding both PpPex3 and PpPex14120, 121. Phosphorylatable variants of the

14 classical AIMs have been reported for both Atg36 and PpAtg30. However, disruption of these

AIMs only delays pexophagy rather than abrogating it145. As previously mentioned, Atg11 is required for pexophagy146. PpAtg37 is an integral peroxisomal membrane protein specifically required for pexophagy in P. pastoris. During pexophagy, PpAtg37 is necessary for phagophore formation, as PpAtg37-null cells fail to recruit PpAtg11 to peroxisomes147. In contrast to yeast, no pexophagy-specific cargo receptor has been described in mammals. Rather, mammalian pexophagy relies on the ubiquitination of peroxisomal proteins and their recognition by

SQSTM1130 and NBR1148. Initially, it was reported that PEX3 overexpression leads to peroxisome ubiquitination and pexophagy induction149. However, blocking PEX3 ubiquitination does not prevent pexophagy and this study did not determine the specific peroxisomal proteins targeted for ubiquitination. Subsequently, two studies identified PEX5 mono-ubiquitination as the cargo signal for peroxisome degradation150, 151. PEX5 is a cytosolic protein that shuttles between the peroxisomal membrane and the cytosol in a ubiquitin-dependent manner152. Accumulation of monoubiquitinated PEX5 on the peroxisomal membrane, which is unable to shuttle back to the cytosol, triggers pexophagy150. Furthermore, in response to reactive oxygen species (ROS), PEX5 is phosphorylated and subsequently mono-ubiquitinated, which leads to pexophagy induction in a

SQSTM1-dependent manner151. A recent study has indicated that the peroxisomal E3-ubiquitin ligase PEX2 is responsible for PEX5 ubiquitination153. These data suggest a model in which mammalian pexophagy is dependent on membrane accumulation of ubiquitinated peroxisomal proteins, such as PEX5, that are recognized by the ubiquitin-binding receptors SQSTM1 and

NBR1, which in turn link the target peroxisomes to LC3/GABARAP-bound sequestration membranes (Figure 1.8). However, this simple model fails to answer several questions. From a mechanistic perspective, how does PEX5 ubiquitination at a specific site determine whether the

15 protein shuttles into the peroxisome or is directed to proteasomal degradation? Are there distinct mechanisms involving ROS and amino-acid-starvation-induced pexophagy? Regarding this last point, other studies have reported that the peroxisomal membrane protein PEX14, which acts as a docking factor for PEX5, can directly interact with LC3-II under starvation conditions, outcompeting PEX5154. This opens the possibility of different pathways being involved under different pexophagy-inducing stimuli. Finally, the human Atg37 ortholog, ACBD5, has also been reported as an essential pexophagy factor147. It will be interesting to determine the role of ACBD5 in pexophagy and its connection to possible undiscovered mammalian pexophagy receptors.

1.5.3 Mitophagy

Mitophagy is a critical quality control process that eliminates damaged and/or superfluous mitochondria through their selective autophagic degradation155, 156. Deficiencies in mitophagy have been linked to development of several pathologies, including neurodegenerative disorders such as Parkinson’s disease157. Mitochondria have multiple metabolic functions and also influence cell fate by regulating apoptosis. Consequently, mitochondrial damage leads to loss of metabolic homeostasis. In addition, disruption of oxidative phosphorylation in damaged mitochondria leads to excessive ROS generation158. Mitochondria are high-maintenance organelles, and non- functioning/superfluous mitochondria become an energetic burden. Therefore, the regulation of mitochondrial quality and quantity is of paramount importance. Although mitochondria harbor some internal quality control machinery159, the major contribution towards maintaining mitochondrial integrity comes from mitophagy, which functions in concert with the UPS to ensure mitochondrial homeostasis160.

In fungi, mitophagy can be triggered by nitrogen starvation161-163 or post-log-phase growth in a non-fermentable medium. In yeast, selectivity is provided by the outer mitochondrial

16 membrane (OMM) receptor Atg32162, which links targeted mitochondria to the autophagic machinery119, 164. The cytosolic N terminus of Atg32 interacts with Atg11165. Ectopic targeting of the Atg32 N terminus to peroxisomes leads to pexophagy, underscoring the function and sufficiency of Atg32 as an autophagy receptor166. The C terminus of Atg32 faces the intermembrane space, and its proteolytic processing by Yme1 may be required for efficient mitophagy156. The interaction between Atg32 and Atg11 promotes recruitment of mitochondria to the PAS for sequestration. Atg32 also orchestrates the subsequent expansion of the phagophore around the mitochondria through its interaction with the cytosolic AIM domain of Atg8155, 166.

However, mutating the Atg32 AIM causes only a partial mitophagy defect, suggesting that the

Atg32–Atg8 interaction increases mitophagy efficiency, but remains auxiliary162, 166, 167.

Expression of Atg32 can be influenced by oxidative stress and nutritional status. In P. pastoris, the Ume6–Sin3–Rpd3 complex, positively regulated by Tor, suppresses ATG32 transcription161. During starvation, Tor is inactivated, promoting the synthesis of Atg32 and starvation-induced mitophagy. However, upregulation of Atg32 expression is not, by itself, sufficient to induce mitophagy. Atg32 is activated by phosphorylation at residues Ser114 and

Ser119 in its cytosolic domain, facilitating its interaction with Atg11167. CK2 (casein kinase 2) has been proposed as the Atg32 Ser114 kinase168, as CK2 phosphorylates Atg32 in vitro but fails to phosphorylate Atg32S114A. Similarly, CK2 temperature-sensitive mutants fail to phosphorylate

Atg32168. However, CK2 is a multitasking kinase and its activation is independent of mitophagy- inducing stimuli162. Therefore, other signaling pathways may contribute to the temporal selectivity of CK2-mediated phosphorylation of Atg32. Two mitogen-activated protein kinase (MAPK) pathways have been implicated in mitophagy regulation in yeast169. Hog1 is a MAPK in the Ssk1–

Pbs2 pathway and Atg32 phosphorylation is suppressed in hog1Δ cells. However, Hog1 does not

17 phosphorylate Atg32 in vitro, suggesting indirect regulation167. The Slt2 (another yeast MAPK) pathway plays a role in mitochondrial recruitment to the PAS169. Although further investigation is required to identify the signaling circuit regulating Atg32 phosphorylation, cooperative expression and activation of Atg32 highlights the multiple levels of regulation involved in mitophagy induction.

As the dimensions of intact mitochondria are larger than that of autophagosomes, sequestration of damaged mitochondria might be facilitated by mitochondrial fission162, 165. In S. cerevisiae, mitochondrial fission is mediated by several factors, including Dnm1 and Fis1165.

Deletion of either DNM1 or FIS1 significantly suppresses mitophagy163, 170. Dnm1 interacts with

Atg11, allowing the former to be recruited to mitochondria targeted for degradation163. The proteins associated with the ER–mitochondrial encounter structure (ERMES) may also play a role in modulating mitochondrial fission during mitophagy165. Nevertheless, the exact mechanism of mitophagy-associated mitochondrial fission is unclear, and as yet unidentified fission factors may be involved.

Mitophagy in mammals is mechanistically more complex than in yeast and is induced by cellular and developmental cues. In mammalian cells, loss of mitochondrial membrane potential is a potent inducer of mitophagy8, 171. However, while the use of chemicals that target the electron transport chain, or act as protonophores, is a convenient and efficient way to study mitophagy, the acute dissipation of mitochondrial membrane potential precludes the study of subtle regulatory phenomena172. Furthermore, such severe mitochondrial damage might not be representative of the true pathophysiological triggers.

In mammals, mitophagy plays important physiological roles in development and cellular differentiation. Erythrocyte development requires selective degradation of mitochondria in

18 reticulocytes173, and embryonic development in some organisms involves selective degradation of paternal mitochondria in the zygote174. Hypoxia, which disrupts mitochondrial respiration, is another stimulus that promotes mitophagy in mammalian cells155.

The PINK1–PRKN/PARK2 pathway is the most extensively characterized mechanism effecting mitochondrial quality control in most mammalian cells. PINK1 is a Ser/Thr kinase with a C-terminal kinase domain and N-terminal mitochondrial targeting sequence155, and

PRKN/PARK2 (also known as parkin) is an E3-ubiquitin ligase175. Loss of mitochondrial integrity is usually accompanied by mitochondrial depolarization. PINK1, which requires the mitochondrial membrane potential for its inner mitochondrial membrane (IMM) import, acts as a depolarization sensor176. In healthy mitochondria, PINK1 is imported into the matrix where it is cleaved by proteases and subsequently released back into the cytosol for degradation through the N-end rule pathway177, 178. In compromised mitochondria, the loss of membrane potential prevents translocation, and PINK1 is stabilized on the OMM, leading to its activation by autophosphorylation172, 176, 179. Active PINK1 phosphorylates several substrates including ubiquitin, MFN1 (mitofusin 1), MFN2 and PRKN/PARK2176, 180. Unphosphorylated

PRKN/PARK2 is autoinhibited176, 180, whereas PINK1-mediated phosphorylation of

PRKN/PARK2176 leads to activation. PINK1 also phosphorylates available ubiquitin attached to

OMM proteins at Ser65, generating phospho-ubiquitin181, 182, which acts as a PRKN/PARK2 substrate177. PRKN/PARK2 subsequently links phosphoubiquitin chains to OMM proteins, which possibly results in a feed-forward amplification loop, recruiting more PRKN/PARK2176.

Phosphorylation of MFN2 by PINK1 might also play a role in PRKN/PARK2 recruitment183, possibly acting along with phospho-ubiquitin at the OMM. However, the role of MFN2 in PRKN/

19 PARK2 recruitment is controversial184. The classic model for mitophagy involves recognition of polyubiquitinated mitochondria by autophagy receptors SQSTM1 and OPTN, which bind LC3176,

185. This interaction tethers damaged mitochondria to the expanding phagophore and promotes their subsequent sequestration within autophagosomes (Figure. 1.9A). Recent progress in the field suggests a complementary model, whereby PINK1-mediated phosphorylation of ubiquitin, independent of PRKN/PARK2 activity, is sufficient to recruit autophagy receptors CALCOCO2

(also known as NDP52) and OPTN and induce low-amplitude mitophagy186. In this model,

CALCOCO2 and OPTN can successfully recruit the Ser/Thr kinase ULK1 and facilitate mitophagy initiation upstream of LC3 binding186. The importance of PRKN/PARK2-mediated ubiquitination is indicated by the fact that overexpression of the mitochondrial deubiquitinase

USP30 inhibits mitophagy by promoting deubiquitination of PRKN/PARK2 substrates187.

Polyubiquitination also acts as a signal that promotes VCP (valosin containing protein, also known as p97)-mediated extraction of OMM proteins and their subsequent proteasomal degradation188, causing disruption of the OMM188. Recent findings suggest that OMM disintegration serves to expose the IMM protein PHB2 (prohibitin 2), which possesses a LIR and functions as a mitophagy receptor174. PHB2 promotes mitophagy in a PINK1-PRKN/PARK2-dependent manner, and the selective removal of paternal mitochondria in Caenorhabditis elegans embryos requires PHB2 function174. The PINK1-PRKN/PARK2-dependent generation of mitochondria-derived vesicles

(MDVs)189, 190 is an alternative pathway to conventional PRKN/PARK2-dependent mitophagy.

Limited and localized mitochondrial damage promotes MDV formation to ensure selective removal of damaged portions of a mitochondrion, instead of the entire organelle191. It is possible that the PINK1-PRKN/PARK2 pathway switches between MDV formation and mitophagy, depending on the extent of mitochondrial damage155.

20 PINK1 and PRKN/PARK2 are also involved in regulating the arrest of mitochondrial motility following mitochondrial damage192. Mitochondria are transported by the kinesin KIF5 on microtubules. KIF5 binds mitochondria through the adaptor TRAK1-TRAK2 and the OMM protein RHOT1 (also known as MIRO1)193. Following mitochondrial damage, RHOT1 is one of the earliest proteins to be degraded by PRKN/PARK2-mediated ubiquitination194, a process that also requires the interaction of RHOT1 with the LRRK2 kinase195. The removal of RHOT1 halts mitochondrial motility and quarantines damaged mitochondria for degradation195. In cells that harbor mutations in PINK1, PRKN/PARK2 or LRRK2, RHOT1 degradation is inhibited, leading to continued motility of damaged mitochondria and delayed mitophagy196.

Not all mammalian cell types express PRKN/PARK2 and several mitochondria-localized mitophagy receptors exist in mammalian cells. BNIP3L (also known as Nix) is one such mitophagy receptor and is involved in the selective elimination of mitochondria during the differentiation of reticulocytes into erythrocytes173, 191. BNIP3L localizes to the OMM and contains a LIR near its cytosolic N terminus194, the activity of which may be regulated by phosphorylation134. However, mutations in the BNIP3L LIR only lead to a partial loss in mitophagy196. Another short motif has recently been reported to be indispensable for BNIP3L function197. Although the exact mechanism by which BNIP3L mediates mitophagy remains unknown, reports suggest that BNIP3L may promote mitochondrial depolarization179, leading to PINK1-PRKN/PARK2 recruitment to mitochondria and activating mitophagy179. BNIP3L might also work in concert with the related protein BNIP3198, which also possesses a LIR199. BNIP3L is also involved in hypoxia-induced mitophagy199, as is the LIR-containing OMM protein, FUNDC1200. Mutations in the FUNDC1

LIR lead to loss of function201. Similar to Atg32, FUNDC1 is regulated by reversible phosphorylation. Under normal conditions, FUNDC1 is phosphorylated by SRC kinase and

21 CSNK2202, including modification of one site in its LIR. Hypoxia promotes dephosphorylation of these residues, involving the phosphatase PGAM5202. Hypoxia-induced mitophagy is particularly relevant to the pathobiology of tumors and elucidating the role of BNIP3L and FUNDC1 in these contexts might be an important step towards therapeutic intervention203, 204.

In mammals, mitochondrial dynamics are regulated by the fission-promoting GTPase

DNM1L (also known as DRP1) and the profusion factors MFN1-MFN2 and OPA1201, 205.

Mitophagy induction is accompanied by a decrease in mitochondrial fusion and an increase in mitochondrial fission to facilitate degradation of damaged mitochondria198. PINK1 activation promotes PRKN/PARK2-mediated degradation of MFN1-MFN2, consistent with the idea of reduced fusion180. The mitophagy receptor FUNDC1 is also involved in regulating mitochondrial dynamics during mitophagy. Whereas FUNDC1 binds to and recruits OPA1 to mitochondria under normal conditions, following mitochondrial damage it preferentially recruits DNM1L, promoting fission206. Similar to ERMES in yeast, mitochondria-associated membranes are sites of ER– mitochondria contact in mammals and have also been proposed to modulate mitophagy-related mitochondrial fission207, although the mechanism remains unclear.

Whereas most selective autophagy receptors are proteins, recent evidence suggests that mitophagy may also be orchestrated by lipid receptors207. Cardiolipin, a lipid unique to mitochondria, may act as a mitophagy receptor in mammalian cortical neurons. Rotenone-induced mitochondrial damage causes a dramatic translocation of cardiolipin from the inner to the outer mitochondrial membrane208, where it interacts with the LC3 N terminus. Inhibition of cardiolipin synthesis or translocation reduces the efficiency of mitophagy in these neurons208. Cardiolipin was also recently reported to modulate mitophagy in S. cerevisiae132 and ceramide has also been

22 implicated as a mitophagy receptor in certain cancer cell lines209.

1.5.4 Reticulophagy

Reticulophagy describes the degradation of the ER by selective autophagy. Perturbation of ER function results in the accumulation of misfolded proteins and ER stress, which in turn triggers the unfolded protein response (UPR) and ER-associated degradation, in order to recover cellular homeostasis210. Autophagy is also activated by ER stress211 as a means to control ER size and counterbalance ER expansion after the UPR212, 213. Other stimuli, such as rapamycin treatment and nutrient starvation, also activate reticulophagy214, 215. Similar to other selective autophagy pathways, cargo receptors have been described for selective ER degradation. In yeast, starvation- induced reticulophagy depends on Atg39 and Atg40—predicted transmembrane proteins that localize to the perinuclear and cytoplasmic ER, respectively. Consistent with their role as cargo receptors, Atg39 and Atg40 contain AIMs, and interact with both Atg8 and Atg11215. In mammals,

RETREG1 (also known as FAM134B) is a reticulophagy cargo-receptor protein, as well as an

Atg40 functional homolog214. Similar to Atg40, RETREG1 localizes to the cytoplasmic ER and interacts with LC3 and GABARAP family members through its LIR (Figure 1.9B). Consistent with the reported role of reticulophagy in controlling ER size, RETREG1 overexpression increases

ER fragmentation, whereas silencing of this protein results in ER expansion.

1.5.5 Nucleophagy

Nucleophagy has been described as the partial or bulk degradation of the nucleus by the vacuole/lysosome. Nucleophagy is closely related to reticulophagy, given that Atg39 localizes to, and mediates the degradation of, the perinuclear ER and nuclear envelope in yeast214. However, to date no Atg39 functional homolog has been described in mammals and it is still unclear how

23 nucleophagy occurs in more complex eukaryotes. However, some studies have suggested selective autophagic degradation of chromatin216 and the nuclear lamina217 could play a role in preventing tumorigenesis.

Other types of nucleophagy, termed piecemeal microautophagy of the nucleus (PMN) or micronucleophagy218, as well as late nucleophagy219 have been described in S. cerevisiae. During

PMN, the outer nuclear envelope protein Nvj1 interacts with the vacuolar membrane protein Vac8.

Together, these forms nuclear–vacuolar junctions that pinch off parts of the nucleus, which are later engulfed and degraded by the vacuole219 (Figure 1.9B). PMN is activated soon after nutrient starvation and depends on the core autophagic machinery220. In contrast, late nucleophagy occurs after prolonged starvation and is independent of Nvj1, Vac8 and some, but not all, core autophagy machinery218. Further studies will be required to understand the individual roles of Atg39-induced nucleophagy, PMN and late nucleophagy during nitrogen starvation.

1.5.6 Lysophagy

Lysophagy is the selective degradation of damaged lysosomes by autophagy. Leakage of lysosomal enzymes into the cytosol, due to lysosomal membrane rupture, leads to lysosomal cell death221. Therefore, removal of damaged lysosomes is necessary to maintain cellular homeostasis.

LGALS3 (galectin 3) binds to glycoproteins exposed by lysosomal membrane damage and colocalizes with LC3, working as a key lysophagy marker221. Even though the specific mechanisms behind lysophagy are yet to be discovered, two independent reports have suggested a model in which damaged lysosomes are selectively degraded in a ubiquitin–SQSTM1–LC3-dependent manner10, 221 (Figure. 1.10A). Thus, lysosome degradation appears analogous to other types of organelle-selective autophagy, such as mitophagy and pexophagy. Still, many questions regarding

24 the specific ubiquitination targets and their regulation remain. Specific physiological conditions in which lysophagy is triggered will need to be determined.

1.5.7 Xenophagy

Xenophagy is the collective term used for selective autophagic degradation of intracellular pathogens, including viruses, bacteria and fungi, which constitutes an important part of the immune response222, 223. Once again, ubiquitination and cargo-receptor binding play an important role in xenophagy. Following Salmonella typhimurium infection and release into the cytosol, bacterial proteins are rapidly ubiquitinated and recognized by the cargo receptors SQSTM1224,

CALCOCO2225 and OPTN225. CALCOCO2 binding to invading bacteria depends on lectin

LGALS8 recruitment to damaged bacteria-containing vesicles226. All three receptors possess ubiquitin-binding domains and LIRs, thus mediating the interaction between ubiquitinated bacteria and LC3/GABARAP family members for phagophore sequestration224, 225, 227 (Figure 1.10B). Wild et al. showed that these three cargo receptors can bind to the same bacterium227. However, individual silencing of SQSTM1, CALCOCO2 or OPTN is sufficient to increase S. typhimurium replication227. This finding suggests that all three cargo receptors have individual roles in xenophagy that cannot be compensated by the other two. Although probably linked to their individual abilities to recruit other autophagy-inducing factors, further studies will be necessary to determine the specific contributions of each cargo receptor. Additionally, finding the specific pathogen proteins that are ubiquitinated will prove indispensable to therapeutically counter the strategies that pathogens have evolved to avoid autophagy.

1.5.8 Lipophagy

Initially discovered in hepatocytes and later in other cell types, lipophagy describes the selective degradation of lipid droplets by autophagy. In vivo and in vitro experiments have shown that

25 lipophagy occurs during basal and starvation conditions, regulating cellular triglyceride content228.

Chaperone-mediated autophagy has been proposed as a regulator of lipophagy. In this model,

CMA would degrade the lipid-droplet-associated PLIN (perilipin) proteins, leading to lipophagy activation3. Although specific receptors for lipophagy have not been found, the metabolic implications associated with this process have highlighted important insights into energy utilization and possible therapeutic strategies for high-fat-diet-induced pathologies. In S. cerevisiae, lipid droplets are degraded in a process termed microlipophagy that depends on the core autophagy machinery, but not Atg11229.

1.5.9 Ferritinophagy

Ferritinophagy involves the degradation of the iron-sequestering protein ferritin230. Iron is an essential component of various enzymes and proteins, making it indispensable for several cellular processes. However, free iron promotes ROS generation and is detrimental to the cell231. Ferritin, consisting of multiple heavy chain (FTH1) and light chain (FTL) subunits, acts as a sink for iron when cellular iron levels are high. Conversely, when bioavailable iron levels are low, ferritin is mobilized by ferritinophagy to release iron230. Ferritinophagy was initially identified in ATG5–/– mouse embryonic fibroblast cells, which fail to degrade ferritin upon iron depletion232. Selectivity during ferritinophagy is mediated by the receptor NCOA4, which specifically binds FTH1 and marks ferritin as autophagic cargo233, 234. The level of NCOA4 is kept low in iron-replete conditions by the iron-dependent interaction between the HECT E3 ligase HERC2 and NCOA4, followed by ubiquitination and proteasomal degradation of NCOA4233. In response to iron depletion, NCOA4 is stabilized, allowing ferritin to be selectively degraded. NCOA4 does not contain a conventional

LIR motif, in contrast to other autophagy receptors230. Therefore, how NCOA4 links its cargo to phagophores promises to be an intriguing question for the field.

1.5.10 Glycophagy

Glycophagy refers to the selective autophagy-mediated degradation of glycogen, the storage form of glucose in animal cells, by GGA (glucosidase alpha, acid) within the lysosome235. Glycophagy is distinct from cytosolic glycogen breakdown by PYG (glycogen phosphorylase); these pathways probably have complementary roles in glycogen catabolism because they preferentially act on slightly different glycogen substrates236. The putative receptor for glycophagy is STBD1 (starch binding domain 1), which possesses a CBN20 glycan-binding domain237 as well as a LIR236.

STBD1 localizes to glycogen particles and binds GABARAPL1235 but not LC3B235. Current evidence indicates an important role for glycophagy in cardiac and hepatic pathophysiology, and further mechanistic investigation of this process will be crucial for realizing the full scope of this pathway in carbohydrate metabolism.

27 Table 1.1 Selective autophagy ligands, receptors and scaffolds in yeast and mammals

Figure 1.1 The 3 main types of autophagy in yeast and mammals A. Yeast cells carry out both macroautophagy and microautophagy. Although macroautophagy consists of the bulk degradation of cytoplasmic material that is sequestered inside double- membrane autophagosomes that then fuse with the vacuole, microautophagy works by directly taking up the substrates through invagination of the vacuole. B. In mammals along with microautophagy and macroautophagy, chaperone-mediated autophagy enables the degradation of specific protein substrates that contain a KFERQ motif that is recognized by chaperones that mediate the translocation of the unfolded protein into the lysosome through LAMP2A.

Figure 1.2 Autophagy induction. A. In yeast in nutrient-rich conditions target of rapamycin complex 1 (TORC1) and protein kinase A (PKA) inhibit autophagy by phosphorylating autophagy related (Atg) 1 and Atg13. During starvation the Atg1 kinase complex is no longer repressed, Atg13 is partially dephosphorylated, and Atg1 is activated. Atg1 then phosphorylates itself and other targets to induce autophagy. B. In mammals in nutrient-rich conditions mechanistic TORC1 (MTORC1) directly binds ULK1 (unc-51 like autophagy activating kinase 1) through RPTOR (regulatory associated protein of MTOR, complex 1) and inhibits ULK1/2 and ATG13 by phosphorylation. Upon starvation MTORC1 dissociates from the ULK1 kinase complex, allowing ATG13 dephosphorylation and activating ULK1/2 that then phosphorylates members of the complex and other targets to induce autophagy.

Figure 1.3 Autophagy regulation. Through STK11/LKB1 (serine/threonine kinase 11), AMP-activated protein kinase (AMPK) senses decreases in the ATP:AMP ratio and phosphorylates TSC1 (tuberous sclerosis 1)-TSC2, which then targets RHEB (Ras homolog enriched in brain), leading to mechanistic target of rapamycin kinase complex 1 (MTORC1) inhibition and autophagy activation. INSR-IGF1R (insulin receptor-insulin-like growth factor 1 receptor) triggers the activation of the class I phosphoinositide 3-kinase (PI3K), inducing the formation of phosphatidylinositol(3,4,5)trisphosphate (PIP3) and AKT/PKB activation; AKT can inhibit TSC1- TSC2, blocking autophagy. PTEN (phosphatase and tensin homolog) works as a PIP3 phosphatase generating phosphatidylinositol(4,5)bisphosphate (PIP2) and inducing autophagy.

Figure 1.4 Class III phosphatidylinositol 3-kinase (PtdIns3K) complexes. Three class III PtdIns3K complexes can be observed in mammals. All of them require PIK3C3/VPS34 (phosphatidylinositol 3-kinase, catalytic subunit type 3), PIK3R4/VPS15 (phosphoinositide-3-kinase, regulatory subunit 4) and BECN1 ( beclin 1, autophagy related). Specific subunits regulate the function of the different complexes. Binding of ATG14 and AMBRA1 (autophagy/beclin-1 regulator 1) leads to autophagy induction. UVRAG (UV radiation resistance associated) and SH3GLB1 (SH3-domain GRB2-like endophilin B1) binding also activates autophagy, whereas binding to RUBCN/RUBICON/KIAA0226 inhibits autophagosome maturation.

Figure 1.5 Autophagosomes have a diverse range of potential membrane sources. The trans-Golgi network, mitochondria, mitochondrial-associated membrane, and endoplasmic reticulum (ER) have been postulated as membrane donors. Omegasomes have been described as the ER structures that work as a platform for autophagosome formation. The phagophore (shown in red) elongates and engulfs part of a cisternae before it buds off the ER and becomes an autophagosome. MAM indicates mitochondria-associated ER membrane.

Figure 1.6 Two ubiquitin-like conjugation systems. Autophagy related (Atg) 8 and Atg12 go through subsequent activation, mediated by Atg7 and conjugation mediated by Atg3 and Atg10, respectively, before covalently binding to phosphatidylethanolamine (PE) in the case of Atg8, and Atg5 in the case of Atg12. Atg8–PE binds both the inner and outer membrane of the autophagosome, but can be deconjugated by Atg4, the same protein that removes the C-terminal arginine initially present at the Atg8 C terminus. Atg12– Atg5 bind Atg16 creating a large multimeric complex that locates to the phagophore and enhances Atg8 lipidation and membrane expansion. GR indicates glycine-arginine.

Figure 1.7 The Cvt pathway and aggrephagy. A. In the yeast Cvt pathway, prApe1, Ape4 and Ams1 are synthesized in the cytoplasm. prApe1 oligomerizes into dodecamers and subsequent higher-order structures that are recognized by the receptor Atg19. Atg19, in turn, binds the scaffold protein Atg11, forming the Cvt complex. Ams1 and Ape4 also oligomerize and bind Atg19. Atg11 brings the Cvt complex to the PAS, where Atg19 binds Atg8–PE, tethering the Cvt complex to the phagophore. B. In both yeast and mammalian aggrephagy, protein aggregates are ubiquitinated (Ub) and subsequently recognized by cargo receptors. In yeast, Cue5 links the ubiquitinated aggregates to Atg8–PE. During mammalian aggrephagy, TOLLIP, SQSTM1, NBR1 and OPTN tether the ubiquitinated aggregates to the phagophore by binding LC3/GABARAP family members. WDFY3 has been described as a scaffold for SQSTM1-dependent degradation.

Figure 1.8 Pexophagy. In S. cerevisiae pexophagy, Atg36 functions as a receptor, linking peroxisomes to the phagophore by binding Pex3 and Atg8–PE. In P. pastoris pexophagy, PpAtg30 acts as a receptor by linking PpPex3 and PpPex14 to PpAtg8–PE. Atg11 functions as a scaffold for both S. cerevisiae and P. pastoris. The current model of mammalian pexophagy involves the E3-ubiquitin ligase PEX2- mediated mono-ubiquitination of PEX5, which in turn is recognized by receptors SQSTM1 and NBR1, tethering peroxisomes to the phagophore. PEX14 has also been reported to link peroxisomes to the phagophore by directly binding LC3 family members.

Figure 1.9 Mitophagy, Reticulophagy and Nucleophagy. A. The yeast mitophagy receptor Atg32 links mitochondria to the phagophore by directly binding Atg8–PE; Atg11 functions as a scaffold. Several cargo receptors (not all shown) have been described for mammalian mitophagy. Mitochondria depolarization leads to PINK1 activation and phosphorylation of ubiquitin and PRKN, and OMM disruption exposes PHB2. Receptors link mitochondria targeted for degradation to the phagophore. B. In yeast reticulophagy, Atg39 and Atg40 have been proposed as receptor proteins. Atg39 mediates degradation of the perinuclear ER, and Atg40 mediates cytoplasmic ER degradation. Both Atg39 and Atg40 link their respective ER sites to Atg8–PE conjugated membranes for sequestration. Atg11 has been proposed as a scaffold protein for both Atg39- and Atg40-mediated reticulophagy. During mammalian reticulophagy, RETREG1/FAM134B tethers the cytoplasmic ER to LC3/GABARAP family members for membrane sequestration and degradation. C. Because Atg39 specifically localizes to the perinuclear ER, Atg39-mediated degradation is also considered to be nucleophagy. During PMN, the nuclear envelope protein Nvj1 and vacuolar membrane protein Vac8 form nuclear– vacuolar junctions, which pinch off and engulf part of the nucleus inside the vacuole.

Figure 1.10 Lysophagy and Xenophagy. A. During lysophagy, unknown lysosomal proteins are ubiquitinated and recognized by SQSTM1, which functions as a receptor, linking the damaged lysosomes with the LC3/GABARAP- conjugated sequestering membranes. LGALS3 binds to exposed lysosomal glycoproteins upon membrane rupture. A specific lysophagy mechanism remains to be elucidated. B. In xenophagy, intracellular pathogens such as viruses and bacteria are recognized and ubiquitinated. SQSTM1, OPTN, CALCOCO2 and NBR1 have been described as receptor proteins.

Chapter 2 The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen Starvation-

Induced Autophagy 2

2.1 Abstract

Macroautophagy/autophagy is a key catabolic recycling pathway that requires fine-tuned regulation to prevent pathologies and preserve homeostasis. Here, we report a new post- transcriptional pathway regulating autophagy involving the Pat1-Lsm (Lsm1 to Lsm7) mRNA- binding complex. Under nitrogen starvation conditions, Pat1-Lsm binds a specific subset of autophagy-related (ATG) transcripts and prevents their 30 to 50 degradation by the exosome complex, leading to ATG mRNA stabilization and accumulation. This process is regulated through

Pat1 dephosphorylation, is necessary for the efficient expression of specific Atg proteins and is required for robust autophagy induction during nitrogen starvation. To the best of our knowledge, this work presents the first example of ATG transcript regulation via 30 binding factors and exosomal degradation.

2 This chapter is reprinted from Damian Gatica, Guowu Hu, Xu Liu, Nannan Zhang, Peter R. Williamson and Daniel J. Klionsky (2018) The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen Starvation-Induced Autophagy. Molecular Cell; 73(2):314-324. doi: https://doi.org/10.1016/j.molcel.2018.11.002.

39 2.2 Introduction

Autophagy is a highly conserved and key catabolic recycling process for cell survival triggered under stress conditions, such as nutrient starvation. During autophagy, a diverse range of cytoplasmic components, including proteins, lipids, and damaged and/or superfluous organelles are sequestered by phagophores that mature into double-membrane vesicles, termed autophagosomes. The autophagosomes subsequently fuse with the vacuole (in yeast and plants) or lysosome (in metazoans), leading to the degradation of the sequestered cargo by resident hydrolytic enzymes. The macromolecules obtained from cargo degradation are then transported back to the cytoplasm for recycling1, 6. During nutrient-rich conditions, autophagy is usually kept at low levels, mostly working as a quality control mechanism238. However, upon starvation, autophagy activity is dramatically increased to compensate for the lack of nutrients. Several autophagy-related (ATG) genes and their corresponding proteins have been characterized based on their function(s) in regulating the various stages of autophagy1, 6. Along these lines, multiple studies have highlighted the importance of the post-translational modification of several Atg proteins13, 239, 240, as well as the transcriptional upregulation of ATG genes upon starvation, to increase autophagy activity240-242. However, less is known about the post-transcriptional regulation of the mRNAs encoding these ATG genes and the mechanisms behind maintaining mRNA stability during autophagy induction. Recently, we reported that following nitrogen starvation, the mRNA decapping enzyme Dcp2 is inhibited, which prevents exonuclease Xrn1-mediated 5’-3’ degradation of multiple ATG transcripts and enables ATG mRNA accumulation and efficient autophagy induction243. Although providing new insights into the post-transcriptional regulation of autophagy, Xrn1-dependent 5’-3’ degradation is not the only RNA decay pathway that has been described. The ribonuclease complex known as the exosome also mediates RNA degradation in a

40 3’to 5’ direction244, 245. While both RNA decay mechanisms have been extensively studied, little is known about the crosstalk between the two pathways. The RNA-binding protein Pat1/Mrt1 has been proposed as a possible link between 5’-3’ and 3’-5’ mRNA degradation246. Pat1 interacts with the heptameric ring-shaped complex formed by the seven Sm-like proteins Lsm1 to Lsm7, which together form the Pat1-Lsm complex247; this complex preferentially binds to the 3’ of oligoadenylated mRNA248, 249. Initially reported as a decapping enhancer, the Pat1-Lsm complex has also been implicated in protecting the 3’ end of oligoadenylated mRNA from shortening by

10–20 nucleotides, also known as trimming250-252. Furthermore, inhibiting the exosome-dependent

3’-5’ mRNA decay pathway leads to the accumulation of trimmed mRNA in pat1Δ and lsm1Δ strains251, providing evidence that the Pat1-Lsm complex can protect the 3’ end of mRNA from exosome-dependent 3’-5’ degradation. Here, we show that upon nitrogen starvation, the Pat1-Lsm complex binds and stabilizes a subset of ATG mRNA by preventing their exosome-mediated 3’-5’ degradation. We provide evidence that this process is regulated through Pat1 dephosphorylation, is necessary for the efficient expression of specific Atg proteins and is required for normal autophagy induction during nitrogen starvation.

2.3 Results

2.3.1 The RNA-binding protein Pat1 is required for normal ATG mRNA accumulation, Atg protein expression, and autophagy induction during nitrogen starvation

To explore the possibility that Pat1 could have a role in regulating autophagy, we generated pat1Δ cells and measured autophagy activity using the quantitative Pho8Δ60 assay253. Briefly, we engineered a mutant Pho8 phosphatase that lacks the first 60 aa of its N terminus, leaving it unable to reach the vacuole through the secretory pathway. However, by inducing nonselective autophagy through starvation, Pho8Δ60 can be delivered to the vacuole, leading to the cleavage of its

41 propeptide by resident hydrolases and the activation of its phosphatase activity. Thus, by measuring phosphatase activity in Pho8Δ60 cells, we can indirectly measure autophagy induction.

A significant decrease in autophagy activity was observed in pat1Δ cells after 3 hr of nitrogen starvation (Figure 2.1A). To extend our analysis, we carried out a GFP-Atg8 processing assay254.

During autophagy induction, Atg8 is covalently conjugated to phosphatidylethanolamine (PE); hence, Atg8-PE can be found on both the inner and outer membrane of the phagophore. When autophagosomes are completed, outer membrane-bound Atg8-PE is deconjugated and recycled back into the cytoplasm; however, inner membrane-bound Atg8-PE is trapped inside the autophagosome, leading to its degradation after vacuolar fusion. Unlike Atg8, GFP is relatively resistant to vacuolar hydrolysis; thus, by tagging GFP to the N terminus of Atg8, we can measure the amount of Atg8 being delivered to the vacuole during autophagy induction. Similar to the

Pho8Δ60 assay, GFP-Atg8 processing showed decreased autophagy induction in pat1Δ cells after

1 and 2 hr of nitrogen starvation, as observed by free GFP levels (Figure 2.1B).

Pat1 interacts with the Lsm1-Lsm7 ring complex, forming the Pat1-Lsm complex247.

Therefore, we decided to measure autophagy in lsm1Δ cells. Similar to pat1Δ cells, LSM1 deletion led to a significant decrease in autophagy activity in both the Pho8Δ60 (Figure 2.2A) and GFP-

Atg8 processing assays (Figure 2.2B). Deleting both PAT1 and LSM1 did not further decrease autophagy activity (Figure 2.2A), suggesting that the effect of Pat1 and Lsm1 on autophagy involves a similar mechanism. Taken together, these data indicate that the Pat1-Lsm complex is necessary for normal autophagy induction during nitrogen starvation.

Pat1 is an RNA-binding protein involved in preventing mRNA degradation251. To determine whether Pat1 plays a similar role with regard to autophagy, we measured ATG mRNA levels by RT-qPCR in pat1Δ cells. Essential autophagy genes ATG1, ATG2, ATG7, and ATG9

42 were selected as autophagy induction markers due to their high transcriptional induction during nitrogen starvation242. In nutrient-replete conditions, pat1Δ cells showed no significant difference relative to the wild-type control (Figure 2.1C). Conversely, after 1 hr of nitrogen starvation, PAT1 deletion caused a significant decrease in ATG1, ATG2, ATG7, and ATG9 transcripts compared to wild-type cells. Similar results were obtained when checking the levels of the same ATG transcript in lsm1Δ cells (Figure 2.2C). These results suggest that the presence of the Pat1-Lsm complex is necessary for normal ATG mRNA stabilization and accumulation during nitrogen starvation. To exclude the possibility that deleting PAT1 could have an effect on ATG mRNA transcription, we constructed a Pat1-inducible degradation strain using the auxin-inducible degron (AID) system255 and measured ATG mRNA levels after 1 hr of nitrogen starvation in the presence or absence of the mRNA synthetic inhibitor 1,10 phenanthroline243. After 30 min of auxin treatment, Pat1 levels decreased dramatically in our Pat1-inducible degradation strain (Figure 2.2D). ATG1, ATG2, and

ATG7 mRNA levels decreased significantly after transcriptional inhibition with 1,10 phenanthroline or following Pat1 degradation using auxin. Importantly, after both transcriptional inhibition and Pat1-induced degradation, ATG1, ATG2, and ATG7 mRNA levels were further decreased compared to 1,10 phenanthroline- or auxin-treated cells. These results indicate that the

PAT1 deletion effect on ATG mRNA levels are not solely due to lower rates of ATG gene transcription, but rather a post-transcriptional regulation of ATG mRNA.

To determine whether the decrease in ATG mRNA levels observed in the PAT1 deletion cells during nitrogen starvation translated to lower Atg protein levels, we measured the amounts of the Atg1, Atg2, Atg7, and Atg9 proteins by western blot. After 2 and 3 hr of nitrogen starvation, the levels of these protein in pat1Δ cells failed to increase to the same extent as in wild-type cells

(Figure 2.1D). Similarly, after nitrogen starvation, Atg1 and Atg9 levels in lsm1Δ cells failed to

43 increase to the same level as in wild-type cells (Figure 2.2B). Together, these data indicate that the

Pat1-Lsm complex is required for normal ATG mRNA accumulation, Atg protein expression, and autophagy induction during nitrogen starvation.

2.3.2 Pat1 stabilizes ATG mRNA during nitrogen starvation by preventing efficient 3’-5’ degradation

Our data suggest that the Pat1-Lsm complex could be stabilizing ATG mRNAs during nitrogen starvation. The Pat1-Lsm complex protects mRNA from 3’ end trimming and 3'-5' degradation by the exosome250-252. Efficient 3' to 5' mRNA degradation by the exosome requires the Ski complex256, 257. The Ski complex, formed by Ski2, Ski3 and Ski8 258, together with the adaptor protein Ski7259, has been proposed to work as a scaffold to modulate the cytoplasmic functions of the exosome without affecting its nuclear role256, 257. Deletion of any members of the Ski complex produce similar phenotypes 256-258 and prevent efficient 3' to 5' mRNA decay251. To test if the decrease in autophagy observed in pat1Δ cells during nitrogen starvation could be rescued by inhibiting the exosome-mediated 3' to 5' mRNA decay, we constructed pat1Δ ski2Δ and pat1Δ ski3Δ double-deletion strains and measured autophagy by the Pho8Δ60 assay.

Both pat1Δ ski2Δ and pat1Δ ski3Δ strains showed a significant increase in autophagy activity after 3 h of nitrogen starvation compared to pat1Δ cells (Figure 2.3A). Consistent with this result, the GFP-Atg8 processing assay also showed that autophagy activity could be recovered in pat1Δ cells by deleting SKI3 (Figure 2.3B). Similar results were obtained when both LSM1 and

SKI3 were deleted (Figure 2.4A and 2.4B). Furthermore, pat1Δ ski3Δ cells showed normal Atg1,

Atg2, Atg7 and Atg9 protein expression after nitrogen starvation (Figure 2.3C) and similar results were seen with lsm1Δ ski3Δ cells compared to lsm1Δ cells (Figure 2.4B). Correspondingly, ATG1,

ATG2, ATG7 and ATG9 mRNA levels after 1 h of nitrogen starvation were significantly higher in

44 the double mutant than in pat1Δ cells and were similar to wild-type levels (Figure 2.3D). Although

ATG9 mRNA levels significantly increased after nitrogen starvation in pat1Δ ski3Δ and ski3Δ cells compared to wild-type cells, this increase did not lead to a robust increase in Atg9 protein levels.

Together, these data indicate that the Pat1-Lsm complex prevents exosome-mediated 3' to 5' ATG mRNA degradation during nitrogen starvation, and this function is necessary for ATG mRNA stabilization, Atg protein expression and normal autophagy induction during starvation.

We decided to extend our analysis on the effect of deleting PAT1 to additional ATG genes by examining pat1Δ cells with RT-qPCR. Several other ATG genes shared the same mRNA accumulation pattern observed for ATG1, ATG2, ATG7 and ATG9, with no significant differences observed in transcript levels during nutrient-replete conditions and significantly lower mRNA levels after nitrogen starvation (Figure 2.4C). However, a subset of ATG transcripts including

ATG4, ATG17 and ATG18 failed to show any significant difference between pat1Δ cells and wild- type cells during both nutrient-replete and nitrogen starvation conditions. Furthermore, ATG3 and

ATG8 mRNAs showed distinct patterns of transcript accumulation in pat1Δ cells. ATG3 mRNA levels were unchanged during nutrient-rich conditions and significantly increased during nitrogen starvation. Conversely, ATG8 transcripts significantly increased during nutrient-replete conditions and remained unchanged compared to wild-type cells after nitrogen starvation. These data suggest

Pat1 is necessary for the accumulation of a specific subset of ATG transcripts.

2.3.3 Pat1 phosphorylation regulates ATG mRNA stabilization, Atg protein expression, and autophagy induction during nitrogen starvation

Pat1 has been reported to be phosphorylated in yeast at residues S456 and S457 during nutrient- replete conditions, but to rapidly lose this phosphorylation after glucose starvation. Accordingly, we decided to evaluate if nitrogen starvation could lead to Pat1 dephosphorylation by analyzing

45 the levels of phosphorylated and dephosphorylated Pat1 using SDS-PAGE and a phosphate- binding tag (Phos-tag); Phos-tag decreases the migration speed of phosphorylated proteins when resolved by SDS-PAGE, thus providing better resolution from their dephosphorylated counterparts. Under growing conditions phosphorylated Pat1 levels were higher than that of dephosphorylated Pat1; however, starting at 30 min of nitrogen starvation, a shift in the pool of phosphorylated Pat1 to its dephosphorylated state could be observed (Figure 2.5A); nitrogen starvation led to a significant decrease in Pat1 dephosphorylation after 60 min and ultimately resulted in a shift to a lower molecular mass form (Figure 2.5A and 2.5B). Site-directed mutagenesis of Pat1 S456 and S457 residues to alanine or glutamate have been effectively used to study the effects of nonphosphorylatable and phosphomimetic mutants of Pat1, respectively 248,

260, 261. We decided to test if Pat1 phosphorylation on these two serine residues could affect autophagy activity by generating Pat1S456,457A (Pat1-AA) and Pat1S456,457E (Pat1-EE) nonphosphorylatable and phosphomimetic mutants, respectively, both containing the PA tag. Pat1-

AA migrated similar to Pat1-PA after 30-60 min of nitrogen starvation (Figure 2.5A), supporting the conclusion that Pat1 was dephosphorylated under these conditions. Whereas Pat1-AA cells failed to show any difference in autophagy activity as measured by both the Pho8Δ60 and GFP-

Atg8 processing assays, Pat1-EE cells showed decreased autophagy levels similar to pat1Δ cells

(Figure 2.6A and 2.6B). This result suggested that Pat1 dephosphorylation is required for normal autophagy induction during nitrogen starvation.

Next, we sought to determine if Pat1 phosphorylation was important for normal Atg protein expression during nitrogen starvation. In agreement with our autophagy activity data, Pat1-AA cells showed no significant difference relative to wild-type cells, whereas Pat1-EE cells showed a significant decrease in Atg1, Atg2, Atg7 and Atg9 protein expression compared to wild-type cells,

46 again similar to pat1Δ cells (Figure 2.6C and 2.6D). Finally, to determine if the effects on Atg protein expression in Pat1-EE cells were due to a decreased stabilization/accumulation of ATG mRNA, we measured ATG transcript levels by RT-qPCR. Under nutrient-replete conditions no significant difference in ATG mRNA could be observed. However, after 1 h of nitrogen starvation,

Pat1-EE cells showed the same significant decrease in ATG mRNA as pat1Δ cells (Figure 2.6E).

Conversely, in Pat1-AA cells only ATG9 showed a significant increase in mRNA levels compared to wild-type cells; however, as shown in figures 3C and 3D, this increase in ATG9 mRNA did not translate into a significant increase in Atg9 protein levels. Neither ATG17 nor ATG18 mRNA, genes for which the transcript levels remained unaffected in pat1Δ cells (Figure 2.4C), showed a significant difference in Pat1-EE and Pat1-AA cells (Figure 2.6E), further indicating that Pat1 and its regulation by phosphorylation only affects a subset of ATG mRNAs. Together, these data indicate that nitrogen starvation leads to Pat1 dephosphorylation, and that dephosphorylation on residues S456 and S457 is required for specific ATG mRNA stabilization, subsequent protein expression and normal autophagy induction.

2.3.4 Pat1 binds specific ATG mRNAs during nitrogen starvation

Various reports have indicated that the Pat1-Lsm complex has the ability to bind to the 3' end of oligoadenylated mRNAs 248, 249, 252, 262, which is proposed to protect the mRNA from exosome- mediated degradation 246, 250, 251. Our data indicate that the Pat1-Lsm complex can prevent the 3' to

5' degradation of certain ATG mRNAs during nitrogen starvation and that this process is regulated by Pat1 phosphorylation on serine residues 456 and 457. To determine if Pat1 binds to ATG mRNA we performed RNA immunoprecipitation (RIP) 263. Briefly, Pat1 protein A (PA)-tagged and - untagged cells were starved for 2 h, followed by crosslinking. Lysates were immunoprecipitated

47 and crosslinking was reversed. After DNAse treatment, RNA was extracted and quantified using

RT-qPCR.

Our RIP analysis showed that Pat1-PA cells had a significant enrichment in ATG1, ATG2,

ATG7, and ATG9 mRNA, but not ATG17 and ATG18 mRNA compared to ALG9 mRNA, which was used as a negative control (Figure 2.7A). This result indicates that Pat1 preferentially binds

ATG1, ATG2, ATG7, and ATG9 mRNA during nitrogen starvation. To further test if Pat1 interaction with ATG mRNA correlated with its effect on ATG mRNA accumulation, we designed a protein-RNA bimolecular fluorescence complementation (protein-RNA BiFC) assay based on the single-molecule analysis of gene expression 264. In brief, an ATG mRNA of interest was tagged with MS2, an RNA hairpin sequence from bacteriophage, without altering the mRNA original 5'

UTR and 3' UTR. The MS2 coat protein (MCP) was fused to the C terminus of the Venus variant of the yellow fluorescent protein, vYFP (VC), generating MCP-VC. Similarly, a protein of interest

(Pat1, Pgk1 or Dhh1) was tagged with the N terminus of vYFP (VN). By tagging MS2 to the ATG mRNA of interest and MCP to VC, we would expect to observe a vYFP fluorescence signal if the

VN-tagged protein of interest interacts with the ATG mRNA of interest (Figure 2.7B).

During nutrient-rich conditions no fluorescent signal was detected in cells expressing Pat1-VN with either ATG1-MS2, ATG2-MS2 or ATG9-MS2 (Figure 2.8A and 2.8B). Conversely, after 2 h of nitrogen starvation, puncta formation could be observed in Pat1-VN cells expressing ATG1-

MS2, ATG2-MS2 or ATG9-MS2 (Figure 2.7C and Figure 2.8B). This result further indicates that

Pat1 binds to these ATG mRNAs during nitrogen starvation. Pgk1-VN cells expressing ATG1-

MS2, ATG2-MS2 or ATG9-MS2, which were used as negative controls, failed to form any bright puncta during both nutrient-replete and nitrogen-starved conditions. Conversely, Dhh1-VN cells expressing ATG1-MS2 or ATG9-MS2, which were used as positive controls, formed puncta during

48 both growing and nitrogen-starved conditions, with the latter inducing the formation of a higher number of puncta (Figure 2.7C and Figure 2.8A). Dhh1 interaction with ATG1 and ATG9 mRNA during growing conditions is consistent with previous data, highlighting the importance of Dhh1 in regulating ATG1 and ATG9 mRNA decapping and degradation when autophagy is not induced

243. Consistently, ATG18 mRNA which failed to show enrichment during our RIP assay, also failed to show Pat1 binding in our protein-RNA BiFC assay, because Pat1-VN cells expressing ATG18-

MS2 did not produce any puncta during either growing or nitrogen-starved conditions (Figure

2.8C). In contrast, our positive control Dhh1-VN cells expressing ATG18-MS2 showed puncta formation after nitrogen starvation (Figure 2.8C). The fact that Dhh1-VN cells expressing ATG18-

MS2 failed to show any puncta during growing conditions, is in line with our previous results showing ATG18 mRNA is not affected by DHH1 deletion 243.

To determine the effects of Pat1 phosphorylation on ATG1 and ATG9 mRNA binding we performed our protein-RNA BiFC assay using Pat1-VN EE and Pat1-VN AA, phosphomimetic and nonphosphorylatable mutants, respectively. In both ATG1-MS2- and ATG9-MS2-expressing cells Pat1-VN AA exhibited a significant increase in the number of cells forming puncta after 2 h of nitrogen starvation. Furthermore, cells expressing Pat1-VN EE showed a significant decrease in the number of cells forming puncta with both ATG1-MS2 and ATG9-MS2 after nitrogen starvation (Figure 2.7C and 2.7D). Together, these data indicate that Pat1 binds a specific subset of ATG mRNA during nitrogen starvation and that this process is controlled by phosphorylation at residues S456 and S457.

49 2.3.6 Preventing efficient 3’-5’ mRNA degradation recovers autophagy activity in Pat1-EE phosphomimetic cells

Both PAT1 deletion and the Pat1-EE phosphomimetic point mutation led to a similar decrease in autophagy activity, ATG mRNA accumulation and Atg protein levels under nitrogen starvation conditions. Because the PAT1 deletion phenotype could be rescued by deleting SKI3, which prevents efficient 3' to 5' mRNA degradation by the exosome, we hypothesized that the Pat1-EE- mediated decrease in autophagy activity during nitrogen starvation could also be prevented by hindering 3' to 5' mRNA degradation. To this end, we constructed Pat1-EE ski2Δ and Pat1-EE ski3Δ cells and measured autophagy activity using the Pho8Δ60 assay. Deletion of either SKI2 or

SKI3 in cells expressing Pat1-EE significantly increased autophagy activity compared to the otherwise wild-type cells (i.e., Pat1-EE) after 3 h of nitrogen starvation (Figure 2.9A). Consistent with this finding, the GFP-Atg8 processing assay also showed that the Pat1-EE-mediated decrease in autophagy activity could be prevented by deleting SKI3 (Figure 2.9B).

We next checked Atg1, Atg2, Atg7 and Atg9 protein levels by western blot during nutrient- rich conditions and after 3 h of nitrogen starvation. As expected, deleting SKI3 in Pat1-EE- expressing cells was able to significantly prevent Atg protein depletion during nitrogen starvation

(Figure 2.9C and 2.9D). Furthermore, Pat1-EE ski3Δ cells showed a significant ATG1, ATG2 and

ATG7 mRNA accumulation after nitrogen starvation compared to Pat1-EE cells. Reminiscent of the pat1Δ ski3Δ effect on ATG9 mRNA levels, Pat1-EE ski3Δ cells showed a significant increase in ATG9 transcript levels compared to wild-type cells, which failed to translate into a significant increase in Atg9 protein levels (Figure 2.9C-E). Once again, ATG17 and ATG18 transcript levels did not show any significant difference among the different strains tested (Figure 2.9E). Together, these data indicate that when Pat1 residues S456 and S457 are phosphorylated, ATG transcripts

50 fail to accumulate due to 3' to 5' mRNA degradation, which in turn leads to lower Atg protein expression and autophagy induction.

2.3.7 Switching the ATG1 3’ UTR with that of ATG17 prevents the PAT1 deletion-mediated decrease in ATG1 mRNA and protein levels

Our data indicate that during nitrogen starvation Pat1 binds to the 3' UTR of a specific subset of

ATG mRNA, such as ATG1, which is necessary for ATG1 transcript stabilization and subsequent protein expression. Because other ATG genes, such as ATG17 and ATG18, were not affected by

PAT1 deletion, we hypothesized that switching the ATG1 3' UTR with that of ATG17 or ATG18 could prevent the decrease in ATG1 mRNA and protein levels observed in pat1Δ cells. To this end, we generated atg1∆ strains in which ATG1 was expressed under the control of it endogenous promoter and contained either the ATG1, ATG7, ATG17 or ATG18 3' UTR. The strain where we replaced the ATG1 3' UTR with that of ATG7 was used as a negative control because ATG7/Atg7 mRNA/protein levels were decreased by PAT1 deletion, similar to ATG1/Atg1. Consistent with our previous results (Figure 2.1), after 3 h of nitrogen starvation atg1Δ pat1Δ cells expressing

ATG1 mRNA with its own 3' UTR showed a significant decrease in Atg1 protein levels compared to atg1Δ cells complemented with ATG1 mRNA having its endogenous 3' UTR (Figure 2.10A and

2.10B). Conversely, after 3 h of nitrogen starvation PAT1 deletion failed to decrease Atg1 protein levels in atg1Δ cells expressing ATG1 mRNA containing the ATG17 3' UTR. In contrast, switching the ATG1 3' UTR with that of ATG7 did not prevent the decrease in Atg1 protein in pat1Δ cells after 3 h of nitrogen starvation.

Similarly, after 1 h of nitrogen starvation, ATG1 mRNA levels, measured by RT-qPCR, were significantly decreased in atg1Δ pat1Δ cells expressing ATG1 mRNA having either the ATG1

51 or ATG7 3' UTR (Figure 2.10C). Conversely, switching the ATG1 3' UTR with that of either

ATG17 or ATG18 prevented the PAT1 deletion-mediated decrease in ATG1 mRNA levels.

Consistent with our previous data (Figure 2.10C), deleting PAT1 still led to a significant decrease in the mRNA levels of ATG2 with its endogenous 3' UTR in these same strains, indicating that suppression of mRNA loss only occurred in cis. These results further indicate Pat1 is necessary for normal ATG1 mRNA accumulation and protein expression during nitrogen starvation and highlight the importance of the ATG1 3' UTR in its post-transcriptional regulation.

2.3.8 The Pat1 mammalian homolog PATL1 is required for normal ATG2 and ATG9 mRNA accumulation and autophagy induction

To determine if the effect of yeast Pat1 on ATG mRNA and autophagy activity is conserved in mammalian cells, we sought to study PATL1, one of the two human homologs of yeast Pat1 265.

The role of PATL1 in mRNA deadenylation and decapping via its interacting protein partners makes it more similar in function to yeast Pat1 in comparison to PATL2, for which a role in mRNA regulation is still not completely understood 266. To determine the effects of PATL1 on ATG mRNA, PATL1 mRNA levels were depleted using siRNA (Figure 2.11A). Similar to our data in yeast, both ATG2 and ATG9 mRNA were significantly reduced after PATL1 depletion, suggesting that PATL1 could play a role in ATG mRNA stabilization in mammalian cells.

Yeast Pat1 was dephosphorylated at serine residues 456 and 457 upon nitrogen starvation

(Figure 2.5A and 2.5B). Previous whole phosphoproteome studies (Hsu et al., PMID: 21659604) had identified two possible phosphorylatable serine residues, S179 and S184 in PATL1 under nutrient-rich conditions, which were confirmed in the present studies by mass spectrometry (Figure

2.11B). To study the effects of S179 and S184 phosphorylation on ATG mRNA levels, plasmids

52 coding for wild-type, phosphomimetic (PATL1S179,184E; PATL1-EE) or unphosphorylatable

(PATL1S179,184A; PATL1-AA) PATL1 were transfected into Jurkat cells, and ATG2 and ATG9A mRNA levels were measured. In nutrient-rich conditions cells expressing PATL1-AA showed a significant increase in ATG9A and ATG2 mRNA levels (Figure 2.11C) compared to cells expressing exogenous wild-type PATL1. Conversely, under the same nutrient-rich conditions

PATL1-EE-expressing cells showed a significant decrease in both ATG2 and ATG9A mRNA levels. Consistent with our previous yeast data, when autophagy was induced through rapamycin treatment, both ATG2 and ATG9A mRNA levels showed a significant increase in PATL1-AA- expressing cells compared to wild-type cells. Conversely, PATL1-EE cells showed a significant decrease in ATG2 and ATG9A mRNA levels under autophagy-inducing conditions. To determine if PATL1 unphosphorylatable or phosphomimetic constructs could affect autophagy activity,

SQSTM1/p62 levels were determined using a flow cytometry-based assay. Briefly, SQSTM1 is a receptor protein that links poly-ubiquitinated substrates to the Atg8 mammalian homolog LC3B.

During autophagy SQSTM1 and the ubiquitinated cargo are sequestered inside autophagosomes and degraded once lysosomal fusion occurs 123. Thus, by measuring SQSTM1 degradation we can monitor autophagy activity. In both nutrient-rich and autophagy-inducing conditions PATL1-AA- expressing cells showed a significant decrease in SQSTM1 levels indicating increased autophagy flux (Figure 2.11D and Figure 2.11E). These results suggest a conserved mechanism in more complex eukaryotes where PATL1 is an important regulatory factor for normal ATG mRNA accumulation and autophagy induction.

2.4 Discussion

Autophagy is a rapid and robust cellular response to cope with nutrient starvation and other types of stress. To this end, rapid induction of ATG mRNA synthesis through transcription factor activation and inhibition of transcriptional and post-transcriptional repressors is essential for quick autophagy induction. Thus, simultaneous induction of ATG mRNA transcription in addition to reducing degradation proves an efficient strategy for triggering robust autophagy induction.

Conversely, during nutrient-rich conditions autophagy is kept at basal levels in part by inhibiting

ATG mRNA synthesis and inducing transcript degradation. In a previous study we described a similar mechanism, in which the decapping enzyme Dcp2 was phosphorylated during nutrient-rich conditions, but rapidly dephosphorylated during starvation leading to its inactivation and the accumulation of ATG mRNA 243. Here we propose an additional mechanism, which furthers our understanding of how ATG mRNA accumulates under conditions of nutrient starvation. In our model, we propose that during nutrient-replete conditions, many of the ATG mRNAs are continuously degraded by Dcp2-dependent decapping and subsequent 5' to 3' Xrn1-mediated degradation 267. This prevents ATG mRNA accumulation and autophagy induction, keeping autophagy at basal levels. However, when cells go through periods of nutrient deprivation, Dcp2- dependent decapping is inhibited, preventing 5' to 3' mRNA degradation. Simultaneously, Pat1 is dephosphorylated leading to Pat1-Lsm complex binding to specific ATG mRNA 3' UTRs, which prevents exosome-mediated 3' to 5' degradation; the result is the accumulation of ATG transcripts and robust autophagy induction (Figure 2.12).

We identified several ATG mRNA targets whose accumulation patterns during nitrogen starvation were affected by PAT1 deletion. Four of these ATG mRNA targets, ATG1, ATG2, ATG7

54 and ATG9 were further examined, and showed increased Pat1 binding after nitrogen starvation.

However, several other ATG mRNAs, including ATG17 and ATG18 transcripts, were not affected by PAT1 deletion and did not show increased Pat1 binding during nitrogen starvation. Further analysis of ATG1 mRNA after replacing its 3' UTR with that of genes not affected by PAT1 deletion, ATG17 or ATG18, further highlighted the importance of the ATG1 3' UTR in Pat1- mediated regulation of ATG mRNA accumulation and subsequent protein expression during nitrogen starvation. While no specific Pat1-Lsm complex mRNA binding sequence has been found, several studies have reported Pat1-Lsm complex affinity for the 3' UTR of oligoadenylated mRNAs carrying U-tracts (6 or more uracils) on or near their 3' end 246, 248, 249, 268. Whereas all ATG mRNAs affected by PAT1 deletion contain uracil repeats near their 3' UTR, other non-affected

ATG transcripts also have uracil repeats close to their 3' end. Thus, U-tracts may be important for

Pat1-Lsm binding, but are not the only required feature. It is possible that the Pat1-Lsm complex binds not a specific sequence, but rather a secondary structure formed in oligoadenylated mRNA containing U-tracts. While this publication was under revision a recent study proposed that the

Pat1-Lsm complex can bind to stress-activated mRNA during hyperosmotic shock 269. Further supporting the idea that the Pat1-Lsm complex can selectively bind and regulate specific mRNAs.

Further studies will be required to determine the exact mechanism by which the Pat1-Lsm complex discriminates between different subsets of ATG and other stress-related mRNAs.

Two specific mRNAs that we evaluated, ATG3 and ATG8, showed distinct patterns when

PAT1 was deleted. ATG8 transcript levels were high during growing conditions and remained similar to the wild-type during nitrogen starvation. In contrast, ATG3 mRNA levels were similar to wild-type in growing conditions and increased during nitrogen starvation. In particular, the

ATG8 mRNA accumulation pattern was similar to the one observed when DHH1 was deleted 243.

55 Pat1 is a key scaffold protein in the recruitment of different components involved in RNA decapping and translation inhibition, including Dhh1 260, 270. Thus, we cannot exclude the possibility that the effects of PAT1 deletion on ATG8 mRNA levels may be an indirect consequence of not recruiting Dhh1. This possibility would explain the similarities in ATG8 transcript accumulation between DHH1 and PAT1 deletion.

PKA has been reported as the kinase responsible for Pat1 phosphorylation on residues

Ser456 and Ser457 260. The same study provided evidence that glucose starvation, a known inactivator of PKA signaling, triggers rapid Pat1 dephosphorylation. Our current study indicates that Pat1 dephosphorylation on these residues also occurs during nitrogen starvation, a condition that inactivates TOR signaling. We observed that nitrogen starvation shifted the pool of phosphorylated Pat1 to its dephosphorylated state, suggesting that the TOR signaling pathway could regulate Pat1 phosphorylation. Furthermore, Pat1 phosphorylation was predicted to decrease following treatment with rapamycin, a well characterized TOR inhibitor 271, and was found to decrease in the mammalian PATL1 (Hsu et al). The roles of both PKA and TOR in negatively regulating autophagy have been extensively studied 272, and possible cross-talk between the TOR and PKA pathways has been proposed 273, 274. Thus, nitrogen starvation could lead to Pat1 dephosphorylation by TOR inactivation and subsequent PKA inhibition.

56 2.5 Materials and Methods

2.5.1 Experimental model and subject detail

All yeast Saccharomyces cerevisiae strains used in this study were constructed in the SEY6210 genetic background (Table 2.1). Deletions and double-deletions were performed using PCR-based methods 275 (Table 2.1). Strains carrying PA-tagged Atg2 and Atg7 maintaining their original 3'

UTRs were generated by PCR-amplifying plasmid pMJ160 and, following transformation, selection and Cre-loxP marker removal as described previously 275. Pat1-PA, and the Pat1S456,457E-

HA and Pat1S456,457A-HA genomic point mutations were generated as previously described 276, 277.

Strains were grown in YPD medium (1% [w:v] yeast extract, 2% [w:v] peptone, and 2% [w:v] glucose) at 30°C to mid-log phase and then collected. Strains grown in YPD medium were shifted into medium lacking nitrogen (SD-N; 0.17% yeast nitrogen base without ammonium sulfate or amino acids, and 2% [w:v] glucose) cultured at 30°C for the indicated time points and then collected. Pho8Δ60 and western blot experiments and analyses were performed as previously described 253, 254. Plasmid pRS-ATG1 (406) expressing ATG1 under its endogenous promoter was used to switch the ATG1 3' UTR with that of ATG7, ATG17 or ATG18 through fast cloning 278.

Strains expressing ATG1 with different 3' UTR sequences were generated by digesting the corresponding plasmids with StuI and transforming in a atg1∆ strain.

Human Jurkat cells were cultured in RPMI 1640 (10% FBS and 1% penicillin and streptomycin) at 37°C in the presence of 5% CO2. Cell transfections were carried out using an

Amaxa Cell line Optimization Nucleofector Kit (LONZA). PATL1 siRNA (E-015591-00-0005) and non-target siRNA pools (D-001919-10-05) were purchased from Dharmacon.

57 2.5.2 RNA and RT-qPCR

The RNA extraction protocol and qPCR primers were published previously 243. cDNA samples were analyzed using a Bio-Rad CFX Connect Real-Time System. Samples were tested in Hard-

Shell® 96-clear well black shell plates (Bio-Rad). The reaction mix (15 μl final volume) consisted of 7.5 μl Radiant™ Green Lo-ROX qPCR kit (Radiant), 0.6 μl each primer (400 nM final concentration), 1.3 μl H2O, and 5 μl of a 1:5 dilution of the cDNA preparation. The thermocycling program consisted of one hold at 95°C for 3 min, followed by 40 cycles of 5 sec at 95°C and 25 sec at 62°C. After completion, a melting curve was generated to verify PCR specificity, as well as the absence of contamination and primer dimers. The transcript abundance in samples was determined using the CFX Manager™ Software regression method. Relative abundance of reference mRNAs and normalization for different total RNA amounts was done as described previously 243.

2.5.3 Auxin-inducible degron (AID) system and transcriptional inhibition

Saccharomyces cerevisiae SEY6210 cells were first transformed with the plasmid pNHK53 (ADH1p-OsTIR1-9MYC). PAT1 was then tagged with AID-9MYC by homologous recombination (see Supplementary Table 2). The DNA fragments used for transformation were amplified with pHIS3-AID*-9MYC (Addgene, 99524; deposited by Dr. Helle Ulrich) as the template DNA. The auxin-inducible degron refers to the 71-116 amino acids of the AT1G04250/ATIAA17 protein in plants. To deplete Pat1 protein levels, the cells were treated with 300 μM 3-indoleacetic acid (auxin; Sigma) or DMSO (vehicle) during mid-log phase growth in YPD medium for 30 min to induce degradation of Pat1. Subsequently samples were collected for western blot. For RNA extraction and RT-qPCR, cells were grown in YPD medium to mid-log phase, washed and shifted

58 to SD-N for nitrogen starvation. Auxin (300 μM), DMSO and/or 1,10 phenathroline (200 μg/ml) was added to the SD-N medium. After 1 h of starvation and treatment, samples were collected as described above.

2.5.4 Protein-RNA bimolecular fluorescence complementation assay

We constructed ATG(X)-MS2 MCP-VC strains through several steps. First, ATG genes of interest were tagged with 24 MS2 hairpins. pDZ415 (24MS2SL-loxP-KANMX6-loxP; Addgene, 45162, deposited by Dr. Robert Singer and Dr. Daniel Zenklusen) was used as a template to PCR amplify the DNA fragments containing 24 MS2 hairpins, homologous regions to the 3' UTR of ATG genes and the KANMX6 marker flanked by loxP sites. Yeast strain SEY6210 was transformed with the

PCR products, and positive colonies were selected for by growth on YPD plates containing G418, then confirmed by PCR. Next, the KANMX6 marker was removed by introducing Cre recombinase into the cells, resulting in ATG(X)-MS2 strains. Next, we made the pCu-MCP-2yeGFP (405) plasmid by two-step cloning. The CUP1 promoter was amplified by PCR and ligated into the pRS405 vector between the XmaI and SpeI sites to make pCu405. Then MCP-2yeGFP together with the CYC1 terminator sequence was PCR amplified from pDZ274 (MET25p-MCP-2yEGFP;

Addgene, 45929, deposited by Dr. Robert Singer and Dr. Daniel Zenklusen) 264 and ligated into pCu405 between the SpeI and SacII sites to make pCu-MCP-2yeGFP(405); this was linearized by

AflII digestion and integrated into the genome of ATG(X)-MS2 strains to make the ATG(X)-MS2

MCP-2yeGFP strains.

Finally, to make the ATG(X)-MS2 CUP1p-MCP-VC strains, we exchanged the 2yeGFP fragment in the genome of the ATG(X)-MS2 CUP1p-MCP-2yeGFP strains with the VC fragment through homologous recombination. To do this, a DNA fragment with the VC peptide-coding sequence and homologous regions from the MCP-coding sequence was amplified by PCR using

59 pFA6a-VC-His3MX6 279 as template. ATG(X)-MS2 CUP1p-MCP-VC strains were generated from

ATG(X)-MS2 CUP1p-MCP-2yeGFP strains by transforming the VC-encoding DNA fragment, selecting colonies and confirming correct integration by PCR. In the background of ATG(X)-MS2

CUP1p-MCP-VC strains, C-terminal tagging of VN at the PAT1, PGK1, and DHH1 loci in the genome was carried out as described previously 279; for primers specifics see Supplementary Table

2. The BiFC strains were examined by fluorescence microscopy under the indicated conditions using softWoRx software (GE Healthcare).

2.5.5 RNA immunoprecipitation

The RNA immunoprecipitation protocol was adapted from a previously published procedure 263.

A Pat1-PA tagged strain and an untagged strain were cultured to mid-log phase and nitrogen starved for 2 h. Cross-linking was performed by adding formaldehyde to 0.8% and shaking slowly for 10 min at room temperature. To stop cross-linking, glycine was added to a final concentration of 0.2 M and cultures were slowly shaken for 5 min. Cultures were then harvested, washed in PBS and resuspended in FA lysis buffer (50 mM HEPES, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1%

Triton X-100, 0.1% sodium deoxycholate, 0.1 % SDS), containing 5 mM PMSF, 1 tablet of complete protease inhibitor cocktail (Roche) and RNasin® PLUS RNAse inhibitor (Promega).

Cultures were vortexed at 4ºC using glass beads to lyse the cells, centrifuged (1000 x g, 1 min) and the supernatant was collected. Samples were sonicated at 4ºC using three 15-sec pulses of 45% amplitude, with a minute break in between when samples were held on ice. Sonicated samples were collected by centrifugation, and the supernatant was collected and then divided into input and

IP fractions. IP fractions were incubated with IgG Sepharose™ 6 Fast Flow beads (GE healthcare

Life Sciences), overnight at 4ºC, while input fractions were frozen in liquid nitrogen and left at -

60 80°C. IP fractions were washed with FA lysis buffer several times, resuspended in RIP elution buffer (50 mM Tris-HCl, pH 7.5, 10 mM EDTA, 1% SDS) and incubated at 70°C for 10 min. IP supernatant and input samples were collected and reverse cross-linking was performed by addition of proteinase K and incubation for 1 h at 42°C, followed by 1 h at 65°C. Next, acid- phenol:chloroform was added to the samples, which were mixed and centrifuged. The aqueous layer of each sample was recovered and 25 μl 3M sodium acetate, 20 μg glycogen, and 625 μl ice- cold 100% ethanol were added to precipitate RNA for 1-2 h at -80°C. Samples were centrifuged, washed with 70% ethanol and dried for 15 min. Pellets were resuspended in 90 μl of RNA-free water after which 10 μl of TURBO Dnase buffer, 2 μl of TURBO™ DNase (TURBO DNA-free™ kit, Invitrogen) and 0.5 μl of RNasin® PLUS RNAse inhibitor were added. Samples were incubated for 45 min at 37°C to eliminate DNA. Following incubation, DNAse was inhibited using the DNAse inactivation reagent that came with the kit. Samples were then subjected to RT-qPCR as described above.

2.5.6 Western blot imaging

Differences in western blot images between panels are due to two different imaging techniques being used. For some images, photographic film was used to developed membranes, whereas imaging was also performed through the ChemiDoc™ Touch imaging system (Bio-Rad).

2.5.7 PATL1 DNA constructs and site-directed mutagenesis

The cDNA encoding full-length human PATL1 was amplified by RT-PCR from total RNA extracted from Jurkat cells using primers PATL1-S-KPN1 and PATL1-A-NOT1 (Table 2.2). and cloned into compatible sites of a pCMV-GFP vector (Addgene; deposited by Connie Cepko).

61 Mutations were introduced by site-directed mutagenesis and all expression constructs were confirmed by DNA sequencing. Primers used to generate expression constructs for the phosphomimic were PATL1-EE1 and PATL1-EE2, and phosphodeficient mutations used PATL1-

AA1 and PATL1-AA2 (Table 2.2).

2.5.8 SQSTM flow cytometry

Jurkat cells were grown in RPMI 1640 media with 10% FBS and 1% penicillin and streptomycin

5 at 37°C with 5% CO2 until cell counts were 3×10 /ml. Cells were treated with rapamycin (200 ng/ml) overnight, and non-treated cells were used as a control. Cells were harvested and washed with 0.2% BSA in PBS twice, and then fixed in fixation buffer (Invitrogen) 1 h at room temperature. After fixation, cells were washed with 1X permeabilization buffer for 30 min at room temperature. Goat serum (10 µl) was added for blocking and incubated for another 30 min at room temperature. SQSTM1/p62 monoclonal antibody (1 µl; Abcam, ab56416) was added, and mouse monoclonal IgG2a (Abcam, ab170191) was used as an isotype control. After incubation for 1 h at room temperature, cells were washed once with 1X permeabilization buffer, and then 1 µl secondary antibody (Alexa Fluor 594 goat anti-mouse) was added. Cells were incubated at room temperature for 1 h and analyzed with a BD Fortessa flow cytometer.

2.5.8 Mass spectrometry and protein identification

Jurkat cells were grown in RPMI, cells were lysed and PATL1 immunoprecipitated using an anti-

PATL1 antibody (Abcam, ab124257); immunoprecipitated proteins were resolved by to SDS-

PAGE. Identification of all proteins was performed on reduced and alkylated, trypsin-digested samples prepared by standard mass spectrometry protocols. The supernatant and two washes (5% formic acid in 50% acetonitrile) of the gel digests were pooled and concentrated by SpeedVac

62 (Labconco) to dryness directly in 200 µl polypropylene auto-sampler vials (Sun Sri). The recovered peptides were resuspended in 5 µl of solvent A (0.1% formic acid, 99.9% water).

Mass spectrometry analysis was performed on an Orbitrap Fusion Tribrid Mass

Spectrometer with in-line chromatography with a PepMap 100 C18 pre-column (3-µm particle size, 75-µm ID, 2-cm length). Nano LC-MS (LC-MS/MS) was performed with a ProXeon Easy- nLC 1000 multi-dimensional liquid chromatograph and temperature controlled Nanospray Flex

Ion Source (ThermoFisher Scientific). Peptides were separated at 200 nl/min using a PepMap 100

C18 column (2-µm particle size, 75-µm ID, 50-cm length). The mobile phase consisted of a linear gradient prepared from solvent A and solvent B (0.1% formic acid, 99.9% acetonitrile) at room temperature. Acquisitions followed a decision tree EThcD/CID methodology with a combination of data-dependent acquisition and an inclusion list 280. The targeted list was formulated to cover all potential m/z for phosphorylated PATL1 peptides and were subjected to fragmentation by

EThcD.

Data processing and databank searching were performed with Peaks Studio 8.5

(Bioinformatics Solution Inc, Waterloo, ON, Canada). The data were searched against the human, pig, and cow proteins deposited in the Uniprot KB (9/2017) and the common Repository of

Adventitious Proteins (theGPM.org). Carbamidomethylation [C] was set as a fixed modification, while oxidation [M], phosphorylation [S/T/Y] and the SILAC labels 13C(6)15N(2) [K] and

13C(6)15N(4) [R] were used as variable modifications. Peptides were filtered at a 0.5% false discovery rate calculated using a decoy sequence approach with a 2 peptides per protein minimum.

63 2.5.9 Quantification and statistical analyses

Microscopy imaging data analysis and image processing was carried out in softWoRx software

(GE Healthcare). Cellular imaging sample sizing were chosen to be the minimum number of independent experiments required for statistically significant results. Western blot images were quantified by ImageJ software. Statistical analyses were performed using GraphPad Prism 6.

Statistical significance was determined in all cases from at least 3 independent experiments using either Student’s t-test or ANOVA. Differences with a P value < 0.05 or lower were considered significant. *p<0.05, **p<0.01, ***p<0.001. Number of independent experiments (n), statistical test utilized, dispersion of measurements and significance is described in the figure legends.

2.6 Acknowledgments

This work was supported by NIH grant GM053396 to DJK and in part by the Intramural Research

Program of the NIH, NIAID to PRW.

64 Table 2.1 S. cerevisiae strains used in this study.

Strain Genotype Reference WLY176 SEY6210 pho13∆ pho8::pho8∆60 253 DGY001 WLY176 pat1∆::LEU2 This study DGY002 DGY001 ski2∆::HIS3 This study DGY003 DGY001 ski3∆::HIS3 This study DGY004 WLY176 ski2∆::HIS3 This study DGY005 WLY176 ski3∆::HIS3 This study JMY347 WLY176 pZEO1-pho8∆60 pCu-GFP-ATG8::LEU2 This study DGY006 WLY176 pZEO1-pho8∆60 pCu-GFP-ATG8::LEU2 pat1::URA3 This study DGY007 DGY006 ski3∆::HIS3 This study DGY008 JMY347 ski3∆::HIS3 This study JMY316 WLY176 ATG2-PA ATG7-PA This study DGY009 JMY316 pat1∆::URA3 This study DGY010 DGY009 ski3∆::HIS3 This study DGY011 JMY316 ski3∆::HIS3 This study DGY012 JMY316 PAT1-HA::HIS3 This study DGY013 JMY316 PAT1-HA S456E S457E::HIS3 This study DGY014 JMY316 PAT1-HA S456A S457A::HIS3 This study DGY015 DGY012 ski3∆::URA3 This study DGY016 DGY013 ski3∆::URA3 This study DGY017 WLY176 PAT1-PA::TRP1 This study DGY018 WLY176 PAT1-PA S456E S457E::TRP1 This study DGY019 WLY176 PAT1-PA S456A S457A::TRP1 This study DGY020 DGY018 ski2∆::HIS3 This study DGY021 DGY018 ski3∆::HIS3 This study DGY022 JMY347 PAT1-PA::TRP1 This study DGY023 JMY347 PAT1-PA S456E S457E::TRP1 This study DGY024 JMY347 PAT1-PA S456A S457A::TRP1 This study DGY025 DGY022 ski3∆::LEU2 This study DGY026 DGY023 ski3∆::LEU2 This study DGY027 WLY176 PAT1-HA::HIS3 This study DGY028 WLY176 PAT1-HA S456E S457E::HIS3 This study DGY029 WLY176 PAT1-HA S456A S457A::HIS3 This study DGY030 DGY027 ski3∆::URA3 This study DGY031 DGY028 ski3∆::URA3 This study DGY032 SEY6210 ATG1-MS2 pCu-MCP-VC::HIS3 PGK1-VN::KAN This study DGY033 SEY6210 ATG1-MS2 pCu-MCP-VC::HIS3 PAT1-VN::KAN This study SEY6210 ATG1-MS2 pCu-MCP-VC::HIS3 PAT1-VN S456E DGY034 S457E::KAN This study

65 SEY6210 ATG1-MS2 pCu-MCP-VC::HIS3 PAT1-VN S456A DGY035 S457A::KAN This study DGY036 SEY6210 ATG1-MS2 pCu-MCP-VC::HIS3 DHH1-VN::KAN This study DGY037 SEY6210 ATG9-MS2 pCu-MCP-VC::HIS3 PGK1-VN::KAN This study DGY038 SEY6210 ATG9-MS2 pCu-MCP-VC::HIS3 PAT1-VN::KAN This study SEY6210 ATG9-MS2 pCu-MCP-VC::HIS3 PAT1-VN S456E DGY039 S457E::KAN This study SEY6210 ATG9-MS2 pCu-MCP-VC::HIS3 PAT1-VN S456A DGY040 S457A::KAN This study DGY041 SEY6210 ATG9-MS2 pCu-MCP-VC::HIS3 DHH1-VN::KAN This study DGY042 SEY6210 ATG2-MS2 pCu-MCP-VC::HIS3 PGK1-VN::KAN This study DGY043 SEY6210 ATG2-MS2 pCu-MCP-VC::HIS3 PAT1-VN::KAN This study DGY044 SEY6210 ATG18-MS2 pCu-MCP-VC::HIS3 DHH1-VN::KAN This study DGY045 SEY6210 ATG18-MS2 pCu-MCP-VC::HIS3 PGK1-VN::KAN This study DGY046 SEY6210 ATG18-MS2 pCu-MCP-VC::HIS3 PAT1-VN::KAN This study DGY047 SEY6210 pNHK53::URA3 PAT1-AID-MYC::HIS3 This study DGY048 SEY6210 atg1∆::HIS3 ATG1-ATG1 3'UTR::URA3 This study DGY049 SEY6210 atg1∆::HIS3 ATG1-ATG7 3'UTR::URA3 This study DGY050 SEY6210 atg1∆::HIS3 ATG1-ATG17 3'UTR::URA3 This study DGY051 SEY6210 atg1∆::HIS3 ATG1-ATG18 3'UTR::URA3 This study DGY052 SEY6210 atg1∆::HIS3 ATG1-ATG1 3'UTR::URA3 pat1∆::LEU2 This study DGY053 SEY6210 atg1∆::HIS3 ATG1-ATG7 3'UTR::URA3 pat1∆::LEU2 This study DGY054 SEY6210 atg1∆::HIS3 ATG1-ATG17 3'UTR::URA3 pat1∆::LEU2 This study DGY055 SEY6210 atg1∆::HIS3 ATG1-ATG18 3'UTR::URA3 pat1∆::LEU2 This study

66 Table 2.2 Primers used in this study

Primer Name Sequence PAT1 deletion For AGCAAAGGTTTTAACCGGAAGTAAGAGCAGCAAGAAGCACTAGCACAGCTGAAGCTTCGTACGC

PAT1 deletion Rev AAAAAAAAATACATGCGTAAGTACATTAAAATTACAGGAAAAATCGCATAGGCCACTAGTGGATCTG

LSM1 deletion For TAAAAGAAAGCAGCCCTCGAATCGAATTAATTCACCAAAACAGCTGAAGCTTCGTACGC LSM1 deletion Rev TACTCCAGGATATATGTTGGTAGTATTGTGTTTTTCTTTCGCATAGGCCACTAGTGGATCTG PAT1 C-terminal GGGTTGGTGTATCGCGATGGTGAAATATCAGAACTAAAGCGGATCCCCGGGTTAATTAA TAG For PAT1 C-terminal AAAATACATGCGTAAGTACATTAAAATTACAGGAAAAATCGAATTCGAGCTCGTTTAAAC TAG Rev PAT1-S456E S457E GCCGCTGCTGTTGCTTCTAAGCAAAGAAGAAGAGAAGAGTACGCGTTCAACAACGGTAAT For PAT1-S456A S457A GCCGCTGCTGTTGCTTCTAAGCAAAGAAGAAGAGCTGCATACGCGTTCAACAACGGTAAT For PAT1-AID-Myc TAG GGGGTTGGTGTATCGCGATGGTGAAATATCAGAACTAAAGCTTCGTACGCTGCAGGTCGA For PAT1-AID-Myc TAG AAAATACATGCGTAAGTACATTAAAATTACAGGAAAAATCCATCGATGAATTCGAGCTCG Rev SKI2 deletion For AACCTAACTCACAAAATTTACTGTACTAATACTAATTTATCAGCTGAAGCTTCGTACGC SKI2 deletion Rev TTTATAAACATGACTCACATTGAGAATAAATGAGCTCTGCATAGGCCACTAGTGGATCTG SKI3 deletion For ACTAAGAACACAGAAAAGAAACACGAAGAGCAGAGGAAATCAGCTGAAGCTTCGTACGC SKI3 deletion Rev TACATTAAGGTTTGATTGACTATCTCGAATCCAAATTTGCATAGGCCACTAGTGGATCTG ATG17 Template- CCTTTTTATTGGGGTTCTTGTGTGATCTGAGATGCAAAAGCG FastCloning For ATG17 Template- ACCGTATCCTTTTTTTCCTTTTTTTTAATTTTGGTGGTTCATCTTCTG FastCloning Rev ATG18 Template- GGCAGCTCTCTTAGCAAAATAATGATCTGAGATGCAAAAGCG FastCloning For ATG18 Template- GCGAGACACTTCCGTGATTAATTTTGGTGGTTCATCTTCTG FastCloning Rev ATG7 Template- ACATTAATTTGGCATTCATATCTAAATGATCTGAGATGCAAAAGCG FastCloning For ATG7 Template- TGTACCAATGCTATTATATGCAAAATATTAATTTTGGTGGTTCATCTTCTG FastCloning Rev ATG17 Insert- CAGAAGATGAACCACCAAAATTAAAAAAAAGGAAAAAAAGGATACGGT FastCloning For ATG17 Insert- CGCTTTTGCATCTCAGATCACACAAGAACCCCAATAAAAAGG FastCloning Rev ATG18 Insert- CAGAAGATGAACCACCAAAATTAATCACGGAAGTGTCTCGC FastCloning For ATG18 Insert- CGCTTTTGCATCTCAGATCATTATTTTGCTAAGAGAGCTGCC FastCloning Rev ATG7 Insert- CAGAAGATGAACCACCAAAATTAATATTTTGCATATAATAGCATTGGTACA FastCloning For ATG7 Insert- CGCTTTTGCATCTCAGATCATTTAGATATGAATGCCAAATTAATGT FastCloning Rev ATG1 deletion For CAGGTTGAAAATATTGAGGCAGAAGATGAACCACCAAAATCAGCTGAAGCTTCGTACGC ATG1 deletion Rev GGTCATTTGTACTTAATAAGAAAACCATATTATGCATCACGCATAGGCCACTAGTGGATCTG ATG1 MS2-TAG For GTTGAAAATATTGAGGCAGAAGATGAACCACCAAAATTAACCGCTCTAGAACTAGTGGAT ATG1 MS2-TAG Rev GGTCATTTGTACTTAATAAGAAAACCATATTATGCATCACGCATAGGCCACTAGTGGATC ATG2 MS2-TAG For AATCAATGATAAGTACAAGTCCAATCGGACTGATTCGTAACCGCTCTAGAACTAGTGGAT ATG2 MS2-TAG Rev ATATGAATTGAATATATATCAAAAATGTCTGCAAAAATTTGCATAGGCCACTAGTGGATC ATG9 MS2-TAG For TGTTAAAGAGTATTACAAGAAGTCTGACGTCGGAAGATAACCGCTCTAGAACTAGTGGAT ATG9 MS2-TAG Rev TATATAGTTATATTGGATGATGTACACGACACAGTCTGCCGCATAGGCCACTAGTGGATC ATG18 MS2-TAG TTGCTTAATATTGTCACAGTATTCCATCTTGATGGATTGACCGCTCTAGAACTAGTGGAT For

67 ATG18 MS2-TAG CGTTGTGACGTACGGAAGGCAGCGCGAGACACTTCCGTGAGCATAGGCCACTAGTGGATC Rev MCP vYFP-C TAG GGTCTCCTAAAAGATGGAAACCCGATTCCCTCAGCAATCGCAGCAAACTCCGGCATCTACCCAGCTGA For AGCTTCGTACGCT MCP vYFP-C TAG ACCATGATTACGCCAAGCGCGCAATTAACCCTCACTAAAGGGAACAAAAGCTGGAGCTCCGAATTCG Rev AGCTCGTTTAAAC DHH1 vYFP-N TAG TATCCTCCACAGCAGGAACATTTCATGGCGATGCCACCTGGTCAGTCACAACCCCAGTATCGGATCCC For CGGGTTAATTAA DHH1 vYFP-N TAG ATTCTTGTTCAAAATCAATAGTAAAAGTATGGTTACAAAGTAATGTAATTCACAATGGAGATTTGGAA Rev TTCGAGCTCGTTTAAAC PGK1 vYFP-N TAG GGAATTATTGGAAGGTAAGGAATTGCCAGGTGTTGCTTTCTTATCCGAAAAGAAACGGATCCCCGGGT For TAATTAA PGK1 vYFP-N TAG GGATGGGGAAAGAGAAAAGAAAAAAATTGATCTATCGATTTCAATTCAATTCAATGAATTCGAGCTC Rev GTTTAAAC PAT1 vYFP-N TAG CAGGTTGAAAATATTGAGGCAGAAGATGAACCACCAAAATCGGATCCCCGGGTTAATTAA For PAT1 vYFP-N TAG GGTCATTTGTACTTAATAAGAAAACCATATTATGCATCACGAATTCGAGCTCGTTTAAAC Rev PATL1-S-KPN1 CGGCGGGGTACCATGTTCCGCTACGAGTCTTTG PATL1-A-NOT1 CGGCGGGCGGCCGCTTATCGTATCCCCTGAACTAGC PATL1-EE1 TCTAACAGGAGGACTGCCAATGATAGGATCAGTATCCCGCCTTGGTAATGCTCGTTCAGAAAG PATL1-EE2 CTTTCTGAACGAGCATTACCAAGGCGGGATACTGATCCTATCATTGGCAGTCCTCCTGTTAGA PATL1-AA1 CTGCCAATGATAGGTGCAGTTGCCCGCCTTGGTAATGCT PATL1-AA2 AGCATTACCAAGGCGGGCAACTGCACCTATCATTGGCAG

Figure 2.1 PAT1 Deletion Lowers Autophagy Activity after Nitrogen starvation by Decreasing ATG mRNA and Protein Levels. A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT) and pat1Δ strains under growing conditions (+N) and after 3 hr of nitrogen starvation (–N). Error bars indicate the standard deviation of four independent experiments. Student’s t test, ***p < 0.001. B. Autophagy was measured by GFP-Atg8 processing in WT and pat1Δ strains under growing conditions and after 1 and 2 hr of nitrogen starvation; a representative image is shown. C. ATG1, ATG2, ATG7, and ATG9 mRNA levels were determined in WT and pat1Δ strains under growing conditions and after 1 hr of nitrogen starvation by RT-qPCR. Error bars indicate the standard deviation of at least six independent experiments. Student’s t test, *p < 0.05 and ***p < 0.001. D. Atg1, Atg2-PA, Atg7-PA, and Atg9 protein levels were measured by western blot in WT and pat1D strains under growing conditions and after 2 and 3 hr of nitrogen starvation; representative images are shown.

Figure 2.2 LSM1 deletion lowers autophagy activity after nitrogen starvation by decreasing ATG mRNA and protein levels. A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT), lsm1∆ and lsm1∆ pat1∆ strains under growing conditions (+N) and following 3 h of nitrogen starvation (–N). Error bars indicate the standard deviation of 4 independent experiments. ANOVA, ***P <0.001. ns, no statistical significance. B. Autophagy was measured by GFP-Atg8 processing in WT and lsm1∆ strains under growing conditions and after 1 and 2 h of nitrogen starvation. Atg1 and Atg9 protein levels were also measured; representative images are shown. C. ATG1, ATG2, ATG7 and ATG9 mRNA levels were determined by RT-qPCR in WT and lsm1∆ strains under growing conditions and following 1 h of nitrogen starvation. D. Pat1 levels were measured by western blot in a Pat1 auxin-inducible degron (AID) strain in nutrient-rich conditions in the presence of DMSO (vehicle)

70 or 300 μM auxin; the upper left panel shows the loss of Pat1-AID-MYC at the 1-h time point. ATG1, ATG2 and ATG7 mRNA levels were quantified by RT-qPCR in Pat1-AID-MYC strains incubated with auxin and/or the synthetic transcriptional inhibitor 1,10 phenanthroline [200 μg/ml], following 1 h of nitrogen starvation. Error bars indicate the standard deviation of 5 independent experiments. Student’s t-test, ANOVA, * P <0.05, ** P <0.01, *** P <0.001.

Figure 2.3 SKI3 deletion prevents the pat1∆-mediated decrease in autophagy. A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT), pat1∆, ski2∆, ski3∆ pat1∆ ski2∆, and pat1∆ ski3∆ strains under growing conditions (+N) and following 3 h of nitrogen

71 starvation (–N). Error bars indicate the standard deviation of 6 independent experiments. ANOVA, **P <0.01 and *** P <0.001. B. Autophagy was measured by GFP-Atg8 processing in WT, pat1∆, ski3∆ and pat1∆ ski3∆ strains under growing conditions and after 2 h of nitrogen starvation; a representative image is shown. C. Atg1, Atg2-PA, Atg7-PA and Atg9 protein levels were measured by western blot in WT, pat1∆, ski3∆ and pat1∆ ski3∆ strains under growing conditions and after 1 or 2 h of nitrogen starvation; representative images are shown. D. ATG1, ATG2, ATG7 and ATG9 mRNA levels were determined in WT, pat1∆, ski3∆ and pat1∆ ski3∆ strains under growing conditions and after 1 h of nitrogen starvation by RT-qPCR. Error bars indicate the standard deviation of 3 independent experiments. ANOVA, ** P<0.01 and *** P <0.001.

Figure. 2.4 SKI3 deletion prevents the lsm1∆-mediated decrease in autophagy. A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT), lsm1∆, ski3∆ and lsm1∆ ski3∆ strains under growing conditions (+N) and following 3 h of nitrogen starvation (–N). Error bars indicate the standard deviation of 3 independent experiments. ANOVA, *P <0.05 and ** P <0.01. B. Autophagy was measured by GFP-Atg8 processing in WT, lsm1∆, ski3∆ and lsm1∆ ski3∆ strains under growing conditions and following 2 h of nitrogen starvation. Atg1 and Atg9 protein levels were also measured; representative images are shown. C. mRNA levels were determined by RT-qPCR in WT and pat1∆ cells for the indicated ATG genes under growing conditions and after 1 h of nitrogen starvation. Error bars indicate the standard deviation of 3 independent experiments. Student’s t-test, * P<0.05.

Figure. 2.5 Pat1 is dephosphorylated under nitrogen-starvation conditions. A. Pat1 phosphorylation (p-) levels were determined in Pat1-PA strains by migration shift in a Phos-tag SDS-PAGE gel under growing conditions (0 min) and following, 30 and 60 min of nitrogen starvation. The Pat1-PA AA strain was used as a positive control. Representative images are shown. B. Quantification of the ratio between dephosphorylated and phosphorylated Pat1-PA under growing conditions and after 60 min of nitrogen starvation. Error bars indicate the standard deviation of 3 independent experiments. Student’s t-test, **P<0.01.

Figure 2.6 Pat1 dephosphorylation on S456 and S457 regulates autophagy by modulating ATG mRNA and protein levels during nitrogen starvation. A. Autophagy activity was measured by the Pho8Δ60 assay in Pat1-PA, pat1∆, Pat1S456,457E-PA (Pat1-PA EE), and Pat1S456,457A-PA (Pat1-PA AA) strains under growing conditions (+N) and after 3 h of nitrogen starvation (–N). Error bars indicate the standard deviation of 5 independent experiments. ANOVA, **P <0.01 and *** P <0.001. B. Autophagy was measured by GFP-Atg8 processing in Pat1-PA, pat1∆, Pat1-PA EE, and Pat1-PA AA strains under growing conditions and after 2 h of nitrogen starvation; a representative image is shown. C. Atg1, Atg2-PA, Atg7-PA and Atg9 protein levels were measured by western blot in Pat1-HA, pat1∆, Pat1-HA EE, and Pat1- HA AA strains under growing conditions and after 3 h of nitrogen starvation; a representative

75 image is shown. D. Quantification of Atg1, Atg2-PA, Atg7-PA and Atg9 protein levels from (C). Error bars indicate the standard deviation of at least 4 independent experiments. ANOVA, *P <0.05, **P <0.01 and *** P <0.001. E. ATG1, ATG2, ATG7, ATG9, ATG17 and ATG18 mRNA levels were analyzed in Pat1-HA, pat1∆, Pat1-HA EE, and Pat1-HA AA strains under growing conditions and after 1 h of nitrogen starvation by RT-qPCR. Error bars indicate the standard deviation of at least 8 independent experiments. ANOVA, *** P<0.001.

Figure 2.7 Pat1 binds specific ATG mRNAs. A. RNA immunoprecipitation was performed in Pat1-PA and wild-type (WT) (untagged) cells. Enrichment (Pat1-PA: WT) of ATG1, ATG2, ATG7, ATG9, ATG17, ATG18 and ALG9 (control)

76 mRNA after 2 h of nitrogen starvation is presented. Error bars indicate the standard deviation of at least 5 independent experiments. ANOVA, * P<0.05, ** P<0.01, *** P<0.001, ns, no statistical significance B. Schematic model for protein-RNA BiFC. Interaction between MS2 coat protein (MCP) tagged with C-terminal vYFP (VC) bound to an MS2 hairpin-tagged ATG mRNA and Pat1-tagged with N-terminal vYFP (VN), leads to a fluorescent signal by a complete vYFP protein. C. Protein-RNA BiFC was used to determine the interaction of Pgk1-VN, Pat1-VN, Pat1-VN AA, Pat1-VN EE and Dhh1-VN with ATG1-MS2- and ATG9-MS2-tagged mRNA after 2 h of nitrogen starvation (–N). D. Quantification of the number of cells showing puncta in Pat1-VN, Pat1-VN EE and Pat1-VN AA cells also expressing either ATG1-MS2- or ATG9-MS2-tagged mRNA. Error bars indicate the standard deviation of at least 3 independent experiments. ANOVA, *P <0.05 and *** P <0.001.

Figure 2.8. Protein-RNA BiFC of ATG2-MS2, ATG17-MS2 and ATG18-MS2. A. Protein-RNA BiFC was used to determine the interaction of Pgk1-VN, Pat1-VN, Pat1-VN AA, Pat1-VN EE and Dhh1-VN with ATG1-MS2- and ATG9-MS2-tagged mRNA during nutrient- replete conditions (+N). Quantification of the percent of cells with puncta is shown on the right. B. Protein-RNA BiFC was used to determine the interaction of Pgk1-VN and Pat1-VN with

78 ATG2-MS2-tagged mRNA during nutrient-replete conditions and following 2 h of nitrogen starvation (–N). C. Protein-RNA BiFC was used to determine the interaction of Pgk1-VN, Pat1- VN and Dhh1-VN with ATG18-MS2-tagged mRNA during nutrient-replete conditions and following 2 h of nitrogen starvation.

Figure 2.9 SKI3 deletion prevents the Pat1S456E- and Pat1S457E-mediated decrease in autophagy. A. Autophagy activity was measured by the Pho8Δ60 assay in Pat1-PA, pat1∆, Pat1-PA EE, Pat1- PA EE ski2∆, and Pat1-PA EE ski3∆ strains under growing conditions (+N) and after 3 h of nitrogen starvation (–N). Error bars indicate the standard deviation of 6 independent experiments. ANOVA, **P <0.01 and *** P <0.001. B. Autophagy was measured by GFP-Atg8 processing in Pat1-PA, Pat1-PA EE, Pat1-PA EE ski3∆ and Pat1-PA ski3∆ strains under growing conditions and after 2 h of nitrogen starvation; a representative image is shown. C. Atg1, Atg2-PA, Atg7-PA

79 and Atg9 protein levels were measured by western blot in Pat1-HA, Pat1-HA EE, Pat1-HA EE ski3∆ and Pat1-HA ski3∆ strains under growing conditions and after 3 h of nitrogen starvation; a representative image is shown. D. Quantification of the data from (C). Error bars indicate the standard deviation of at least 4 independent experiments. ANOVA, *P <0.05, **P <0.01 and *** P <0.001. E. ATG1, ATG2, ATG7, ATG9, ATG17 and ATG18 mRNA levels were analyzed in Pat1-HA, Pat1-HA EE, Pat1-HA EE ski3∆ and Pat1-HA ski3∆ strains under growing conditions and after 1 h of nitrogen starvation by RT-qPCR. Error bars indicate the standard deviation of at least 8 independent experiments. ANOVA, *** P <0.001.

Figure 2.10 Switching the ATG1 mRNA 3' UTR prevents the PAT1 deletion-mediated decrease in ATG1 mRNA and protein levels. A. Atg1 protein levels were measured by western blot in atg1Δ cells and atg1Δ pat1Δ cells expressing ATG1 under its endogenous promoter and with either the ATG1, ATG7 or ATG17 3' UTR during growing conditions and after 3 h of nitrogen starvation; representative images are shown. B. Quantification of the data from (A). Error bars indicate the standard deviation of at least 4 independent experiments. ANOVA, **P <0.01 and *** P <0.001, ns, no statistical significance. C. ATG1 and ATG2 mRNA levels were analyzed by RT-qPCR in atg1Δ and atg1Δ pat1Δ cells expressing ATG1 under its endogenous promoter and with either the ATG1, ATG7, ATG17 or

81 ATG18 3' UTR, after 1 h of nitrogen starvation. Error bars indicate the standard deviation of at least 6 independent experiments. ANOVA, **P <0.01, *** P <0.001, ns, no statistical significance.

Figure 2.11. PATL1 levels and phosphorylation state regulate ATG2 and ATG9A mRNA levels and autophagy activity. A. PATL1, ATG2 and ATG9A mRNA levels were measured in Jurkat cells after non-target or PATL1 siRNA treatment. B. Cell lysate from actively growing Jurkat cells was subjected to PATL1 immunoprecipitation followed by mass spectrometry. Identified phosphorylated residues are indicated. C. Jurkat cells were transfected with constructs expressing PATL1 wild-type, AA, EE or empty vector and ATG2 and ATG9A mRNA levels were determined after control and

83 rapamycin treatment. D. Jurkat cells were transfected with constructs expressing PATL1 wild- type, AA or EE and SQSTM1 protein levels were measured through flow cytometry. E. Quantification of the data from (D). Error bars indicate the standard deviation of at least 3 independent experiments. Student’s t-test, ANOVA, *P <0.05, **P <0.01, *** P <0.001.

Figure 2.12 Pat1 stabilizes a subset of ATG mRNAs during nitrogen starvation-induced autophagy by preventing 3'-5' mRNA degradation.

84 Under normal growth conditions Dcp2 is phosphorylated, leading to ATG transcript decapping and subsequent 5' to 3' degradation by Xrn1 243, 281. This process occurs in parallel with ATG gene transcription inhibition by inactivation of several transcription factors (not shown). Pat1 is phosphorylated on residues Ser456 and Ser457. Nutrient deficiency leads to Dcp2 and Pat1 dephosphorylation. The former prevents decapping and Xrn1-mediated 5' to 3' degradation of ATG mRNAs, whereas the latter leads to Pat1-Lsm binding to ATG transcripts and prevents their exosome-mediated 3' to 5' degradation. This process, in concert with the transcriptional activation of ATG mRNA synthesis, facilitates ATG transcript accumulation and autophagy induction.

85 Chapter 3 The carboxy terminus of yeast Atg13 binds phospholipid membrane via motifs

that overlap with the Vac8-interacting domain3

3.1 Abstract

Macroautophagy/autophagy is a conserved catabolic recycling pathway involving the sequestration of cytoplasmic components inside double-membrane vesicles termed autophagosomes. The autophagy-related (Atg) protein Atg13 is a key member of the autophagy initiation complex. The Atg13 C terminus is predicted to be an intrinsically disordered region

(IDR) harboring a binding site for the vacuolar membrane protein Vac8. Recent reports suggest

Atg13 acts as a hub to assemble the initiation complex, as well as participating in membrane recognition. Here we show that the Atg13 C terminus directly binds to lipid membranes via electrostatic interactions between positively charged residues in Atg13 and negatively charged phospholipids. We identified two sets of residues in the Atg13 IDR that affect its phospholipid- binding properties; these residues overlap with the Vac8-binding domain of Atg13. Our data indicate that Atg13 binding to phospholipids and Vac8 is mutually exclusive, and both are required for efficient autophagy.

3 A modified version of this chapter has been submitted for publication.

86 3.2 Introduction

Autophagy is a catabolic process marked by the formation of the phagophore, a transient cup- shaped structure that matures into a double-membrane organelle called the autophagosome. In yeast, the autophagosome forms at the PAS, where Atg proteins, donor membranes, and cytoplasmic cargo coalesce in preparation for sequestration of superfluous and aberrant cellular materials. The process of autophagy can be subdivided into four ordered steps: initiation/nucleation, elongation/closure, docking/fusion, and degradation282.

The Atg1 kinase complex, comprised of Atg1, Atg13, and the Atg17-Atg31-Atg29 subcomplex, is a fundamental unit for autophagy initiation. This complex promotes autophagosome formation through post-translational regulation of downstream core autophagy proteins283, 284 and by providing the structural backbone for the growing phagophore285. In recent years, multiple structural and biochemical studies involving the Atg1 kinase complex revealed its highly dynamic self-assembling propensity that paints a comprehensive model for recognizing and integrating Atg9-positive vesicles (donor membranes) at the PAS286-289.

In the center of all the interacting proteins is Atg13, a regulatory protein of the Atg1 kinase complex. In nutrient-rich conditions, Atg13 is hyperphosphorylated in a Tor-dependent manner, whereas nutrient depletion leads to Tor inhibition, Atg13 dephosphorylation, and the activation of

Atg1290. Atg13 contains an N-terminal HORMA domain, which has been suggested to recruit

Atg14,291, 292 and a C-terminal domain predicted to be an intrinsically disordered region (IDR) that forms an unstable structure288 harboring multiple protein binding sites. Specifically, the Atg13

IDR was reported to contain at least two Atg17 binding motifs: the 17LR motif (359-389) and

87 17BR motif (424-436)288, an MIT-interacting motif (MIM) that binds to Atg1 (460-521),293 and the C-terminal (567-738) region occupied by the vacuolar peripheral membrane protein Vac8294.

Furthermore, recent in vitro experiments by Rao et al. showed that recombinant purified Atg13 from S. cerevisiae directly binds yeast polar lipids, suggesting that Atg13 acts as a molecular hub to mediate the intricate spatiotemporal organization necessary for Atg1 kinase complex assembly as well as membrane recognition.

Some insights have been gained into assembly of the Atg1 complex283, 285, 295, but the molecular mechanism of yeast Atg13 lipid binding capability remains to be discovered. In contrast, lipid binding by human ATG13 is mediated by a cluster of lysine residues, at the N terminus of the HORMA domain, which are responsible for electrostatic interactions with negatively charged phospholipids. These membrane-binding lysines are evolutionarily conserved in mouse and zebrafish, but not in yeast66, suggesting that a membrane-binding motif of yeast Atg13 is located in a different region than the N terminus of the HORMA domain.

Here we show that the C terminus of yeast Atg13 directly binds to liposomes via electrostatic interactions between positively charged residues in Atg13 and negatively charged phospholipids.

We identified two sets of positively charged residues in the IDR of Atg13 from S. cerevisiae that affect the phospholipid binding properties of Atg13 when mutagenized. These residues reside in short phenylalanine-containing motifs within the amino acid sequence that overlaps with the Vac8- binding domain of Atg13. Our data indicate that the phospholipid- and Vac8-binding of Atg13 are mutually exclusive and both are required for efficient autophagy.

88 3.3 Results

3.3.1 Atg13[571-700] is intrinsically disordered

The C-terminal tail of Atg13, spanning residues 269-738, is predicted to be intrinsically disordered. Several regions of this disordered domain bind proteins including Atg17, Atg1 and

Vac8. In addition to supporting protein-protein interactions, disordered regions bind liposomes and, in some cases, even alter membrane curvature296. As such, we wondered if the previously observed membrane-binding behavior of Atg13 was carried out by some portion of the predicted disordered C terminus of Atg13. After screening several C-terminal constructs for expression and solubility in E. coli, we succeeded in purifying Atg13[571-700]. As predicted, this region was intrinsically disordered as observed by 1D 1H NMR spectroscopy and circular dichroism (CD)

(Figure 3.1 and Figure 3.2).

3.3.2 Atg13[571-700] binds negatively charged liposomes

To test whether Atg13[571-700] interacts with membranes, we performed liposome sedimentation assays with Folch-derived unilamellar vesicles. In liposome sedimentation assays, protein and liposomes are incubated together and subsequently subjected to centrifugation. Due to their size, liposomes will pellet during centrifugation. In the absence of liposomes or if the protein does not interact with liposomes, the protein will remain in the supernatant fraction after centrifugation.

However, if the protein interacts with membranes it will pellet along with the liposomes. Folch vesicles were generated with diameters of 1.0, 0.6, 0.5, and 0.4 µm, incubated with Atg13[571-

700] and subjected to centrifugation. Atg13[571-700] appeared in the supernatant fraction in the

89 absence of liposomes but pelleted when incubated with all sizes of liposome tested (Figure 3.3A and Figure 3.3B). This finding demonstrates that Atg13[571-700] interacts with Folch liposomes and appears to have little specificity for the size of the vesicle.

Full-length Atg13 binds yeast polar lipids, suggesting that electrostatic interactions may play an important role in liposome binding by this protein. To test if this is the case for Atg13[571-

700], we performed liposome sedimentation assays in the presence of increasing concentrations of

NaCl (Figure 3.3 C and Figure 3.3D). If electrostatic interactions are required for membrane binding, then increasing NaCl concentrations will disrupt this interaction. We found that Atg13 bound to Folch liposomes in buffer containing either 50 or 150 mM NaCl, but that binding was almost completely abolished in buffer containing 300 or 500 mM NaCl. This result suggests that

Atg13 binding to liposomes is mediated primarily through electrostatic interactions.

Intrinsically disordered proteins can bind to membranes using hydrophobic amino acids that fold to form an amphipathic helix upon membrane binding297. While our liposome sedimentation data suggest that this is not the case for Atg13[571-700] we wanted to further confirm this by monitoring the secondary structure of Atg13[571-700] using CD in the absence and presence of liposomes. We did not observe any increase in helix formation upon liposome binding by Atg13[571-700], further supporting the idea that electrostatic interactions are the main mechanism of liposome binding (Figure. 3.2).

We next sought to determine which phospholipids are required for Atg13[571-700] binding. During autophagy, PtdIns in the expanding phagophore membrane is phosphorylated by the PtdIns3K complex to generate PtdIns3P. As such, Atg13 may bind PtdIns3P as a means to

90 recognize the phagophore membrane. To determine if Atg13 is able to bind PtdIns3P in liposomes, we generated liposomes containing 76% phosphatidylcholine (PC), 19% phosphatidylserine (PS) and 5% of either PtdIns or PtdIns3P. This concentration of PtdIns3P was chosen as it is frequently used to investigate the interaction of PtdIns3P-binding proteins, including FYVE and PX domains, with liposomes. Liposome sedimentation assays revealed that Atg13[571-700] did not bind to vesicles containing 5% of either PtdIns or PtdIns3P suggesting that the interaction of Atg13[571-

700] with liposomes is not specific for PtdIns3P (Figure 3.3E).

To determine if Atg13[571-700] recognized liposomes based solely on their negative charge we generated liposomes containing PC, PS, PtdIns, and PE, which are the main phospholipids in yeast polar extracts, and cholesterol. Liposomes were generated with 53.25% PC and 19% PS or 29.75% PC and 42.50% PS while keeping the concentrations of PtdIns and PE constant. Atg13[571-700] did not bind to liposomes containing 19% PS but did bind to liposomes containing 42.5% PS (Figure 3.3F and Figure 3.3G). In addition, we tested if PtdIns3P would alter the binding of Atg13[571-700] in these vesicles. Substituting PtdIns3P for PtdIns in these vesicles led to enhanced binding of Atg13. Because Atg13 was unable to bind liposomes containing only

5% PtdIns3P, the increased binding of Atg13[571-700] to PtdIns3P-containing liposomes with PS is likely due to the increased negative charge rather than a specific interaction with PtdIns3P.

Taken together these data suggest that Atg13[571-700] binds negatively charged lipids.

3.3.3 Atg13[571-700] contains 2 distinct liposome binding regions

Due to the requirement of negatively charged phospholipids for Atg13[571-700] binding, we hypothesized that liposome binding by Atg13[571-700] would be mediated by positively charged

91 amino acids. To elucidate the region of Atg13[571-700] that is responsible for liposome binding, we generated a series of truncations, which were designed to remove different positively charged regions from Atg13[571-700]. These constructs included 1) 571-682, which lacks the highly conserved 683KFHK motif; 2) 571-640, which lacks three distinct positively charged regions; and

3) 647-700 which contains the 683KFHK motif, but lacks the positively charged regions spanning

578-584, 610-613, and 640-642 (Figure. 3.4A). Liposome sedimentation assays were performed using these constructs in 50, 150, 300 and 500 mM NaCl. The only construct that had reduced liposome binding at low NaCl concentrations was Atg13[571-640], suggesting that the 641-700 region is important for liposome binding (Figure. 3.5A). In addition, Atg13[571-682] and [647-

700] displayed nearly identical binding to Atg13[571-700] further supporting a role for the 641-

700 region in membrane binding.

To further examine the role of the 683KFHK motif in liposome binding, we mutated this sequence to K683E H685E K686E. Liposome sedimentation assays were performed in 50, 150,

300 and 500 mM NaCl. The K683, H685, K686E mutation led to a 50% reduction in liposome binding, confirming the importance of this region (Figure 3.5B). This result, taken together with the fact that Atg13[571-682] also bound to liposomes, suggests that Atg13[571-700] may contain a second liposome binding region. One possible liposome-binding region spans residues 640-645, which includes both positively charged and hydrophobic residues in the sequence 640KFKSSI. To test whether residues 640-645 are involved in liposome binding we generated three different mutations including 1) F641E I645E, 2) K640A K642A and 3) K640A F641E K642A I645E. The

F641 I645E mutation led to a reduction in liposome binding at all NaCl concentrations tested.

92 However, the K640A K642A and K640A F641E K642A I645E mutations only led to a reduction in liposome binding at 150 mM NaCl. A summary of all truncation and mutagenesis data is summarized in Figure 3.6. These data suggest that Atg13[571-700] contains two distinct liposome- binding motifs, 640KFK and 683KFHK (Figure 3.4A).

3.3.4 Lipid binding motifs in Atg13 are required for efficient autophagy

To determine if nonselective autophagy activity is affected by a chromosomal mutation in the phospholipid binding motifs of the full-length Atg13 protein, we performed the quantitative

Pho8Δ60 assay. Cells with wild-type Atg13 exhibited a substantial increase in Pho8Δ60 activity after 3 h of nitrogen starvation (Figure 3.7A). In comparison, the F641E I645E and K683E H685E

K686E mutants showed an ~30% decrease in Pho8Δ60 activity; the F641G I645G mutant was even more defective, as it exhibited an ~50% decrease in autophagy activity under the same experimental conditions.

To further test the nonselective autophagy activity of Atg13 mutants, we used the GFP-

Atg8 processing assay. In wild-type cells, a strong band corresponding to free GFP could be observed by western blot after 1 h of nitrogen starvation (Figure 3.7B). In contrast, the chromosomal mutation F641E I645E and K683E H685E K686E in full-length Atg13 rendered the yeast cells significantly defective in autophagy flux, as the level of free GFP for these mutants decreased by 40-50%. Combining five mutagenic replacements into a quintuple glutamate mutant did not lead to any further decrease in autophagy activity (Figure 3.4B). The GFP-Atg8 processing

93 assay was far more sensitive to the F641G I645G mutation and revealed an ~75% decrease in the level of free GFP (Figure 3.7B).

To test the autophagy activity of the Atg13 mutants in the absence of its Vac8 binding partner, we expressed these mutants in a vac8Δ strain and measured prApe1 processing (Figure

3.4C and Figure 3.4D). Briefly, in this background, Cvt pathway is blocked due to VAC8 deletion and prApe1 can only be delivered to the vacuole through autophagy. Without Vac8, Atg13F641,I645E was completely unable to transport prApe1 to the vacuole, whereas Atg13K683,H685,686E functioned similar to the wild-type protein (Figure 3.4C and Figure 3.4D). This result reveals that, in the absence of Vac8, the cellular function of the first phospholipid-binding motif (640KFK) was more essential than that of the second motif (683KFHK). However, in the presence of Vac8 (Figure 3.7), the F641G F645G mutant exhibited the strongest autophagy-defective phenotype, suggesting a role of Vac8 in the interaction between the Atg13 C terminus and phospholipid membranes.

3.3.5 Two phospholipid-binding motifs of Atg13 are within the Vac8-binding domain

In yeast, Vac8 interacts with the Atg13 C-terminal sequence, which does not have homology to human ATG13294, 298. A segment of the Atg13 C-terminal region has homology with Nvj1, a nuclear membrane protein that also interacts with Vac8298. The Vac8-Nvj1interaction forms the nucleus-vacuole junction that is essential for piecemeal microautophagy of the nucleus299,220. Vac8 interaction with Nvj1 or Atg13 is exclusive, and competitive, and relies on positively charged residues in the Vac8 groove298. The recently solved crystal structure of Atg13 in the complex with

94 Vac8 revealed that Atg13 employs a relatively long region to interact with Vac8. In the region of

Atg13 that was used for crystallization (residues 567-695), the segment S660-H685 is stabilized in the Vac8 groove (Figure 3.4A) and reveals the Vac8-Atg13 interface, the N- terminus of which interacts with less strength than the C terminus. The phospholipid-binding motifs identified in our study are within the region 571-700, a very similar region to that used in the Vac8-Atg13 crystals.

When compared to the Atg13-Vac8 crystal structure, the first phospholipid-binding motif of Atg13

(640KFK) is near, but outside, the Atg13-Vac8 interface. The second phospholipid-binding motif

(683KFHK) is at the very end of the interface (Figure 3.4A).

To investigate whether mutations in the phospholipid-binding motifs of Atg13 (640KFK and 683KFHK) affect binding with Vac8, we overexpressed a 6xHis-tagged Atg13[571-700] along with Vac8 in yeast multiple-knockout (MKO)300 cells and tested the presence of Vac8 after affinity isolation of Atg13[571-700] using Ni-NTA agarose. Empty vector with a 6xHis-tagged nonspecific (scrambled) sequence of 14 amino acid residues was used as a negative control. The affinity-isolation experiment showed that 6xHis-Atg13[571-700] interacted with Vac8 (Figure

3.8A). This interaction was reinforced by Glu mutations (F641E I645E or K683E H685E K686E) in either lipid binding motif, as more Vac8 was pulled down along with the mutated Atg13 relative to the wild-type under the same experimental conditions (Figure 3.8A). Given that cationic residues in the Vac8 groove are responsible for interaction with Atg13298, glutamate residues of mutants engaged in nonspecific electrostatic interactions could possibly explain this result. In contrast, the F641G I645G mutant could not be detected in the affinity-isolation experiment

(Figure 3.8A), although it was expressed in the cell (Figure 3.9A and Figure 3.9B).

95 One explanation for the absence of Atg13[571-700]F641,I645G in the affinity isolation analysis could be a relatively high susceptibility of this mutant to proteolytic digestion after lysis.

Attaching a nonspecific scrambled sequence to the C terminus of the 6xHis-Atg13[571-

700]F641,I645G mutant improved its stability in the in vitro experiment and yielded a near wild-type level of Vac8 pulled down by Atg13 (Figure 3.8B). The same scrambled sequence also stabilized wild-type 6xHis-Atg13[571-700] (Figure 3.8B). The amount of Vac8 pulled down along with the glutamate mutants (F641E I645E and K683E H685E K686E), which also carried the nonspecific

14 amino acid sequence at the C terminus, was still higher than the level of Vac8 pulled down by wild-type Atg13[571-700] (Figure 3.8B). The 6xHis-Atg13[571-700]D651,Q666,P667R mutant was hardly detectable in the input, and highly inefficient in pulling down Vac8 as compared to the wild-type, despite the presence of the scrambled sequence at its C terminus (Figure. 3.8B). This mutant also displayed very low stability in the total cell lysate (Figure 3.9C). This result shows that the scrambled sequence alone cannot engage in efficient binding to Vac8 unless it is accompanied by an Atg13-specific sequence that is capable of attaching to the Vac8 groove.

Clearly, repulsive forces between arginine residues in the triple mutant and positively charged residues in Vac8 prevented the mutated Atg13[571-700] from binding efficiently to Vac8. As a result, Atg13[571-700] was largely degraded in starved cells. Because mutations of Atg13[571-

700] with or without the scrambled sequence clearly modulated its stability in the cell and the level of Vac8 in the affinity isolation experiments (Figure 3.8A,B, Figure 3.9A-C), we looked at the levels of 6xHis-Atg13[571-700] attached to the scrambled sequence in the MKO cells with or without overexpressed Vac8. We found that the level of 6xHis-Atg13[571-700] linked to the

96 scrambled sequence positively correlated with the level of Vac8 in the cell (Figure 3.9D), a result that further supports our finding that Atg13[571-700] gains cellular stability via interaction with

Vac8. Glutamate mutations in both phospholipid-binding motifs of Atg13 made the interaction with Vac8 more efficient, relative to wild-type. This finding suggests that in the cell both motifs are in proximity to Vac8. Stabilization of 6xHis-Atg13[571-700] by a nonspecific interaction of the C-terminal scrambled sequence with Vac8 indicates that Ser700 in Atg13 is also in proximity to Vac8, in agreement with the Atg13-Vac8 crystal structure.

To test the binding affinity of the Atg13 glutamate mutants with Vac8 relative to wild-type, we applied isothermal calorimetry (ITC) to measure dissociation constants for Atg13[571-700] binding to recombinant purified Vac8[10-515]. The wild-type and mutant Atg13[571-700] proteins all bound to Vac8 with similar affinities, Kd of 680-971 nM (Table 2.2), further confirming that the Atg13[571-700] region carries the Vac8-binding domain, and that the Atg13 residues required for liposome binding are not required for Vac8 binding. These results suggest that the

Vac8-binding region and liposome-binding region are overlapping, but both require distinct residues. This observation opens the possibility that the interaction of Atg13[571-700] with Vac8 and phospholipid membrane is mutually exclusive.

3.3.6 Atg13[571-700] exclusively interacts with either phospholipid membranes or Vac8

If Atg13[571-700] interacts exclusively with either phospholipid membranes or Vac8, the lipid- binding motifs should be masked upon Vac8 binding, and the liposome binding of Atg13[571-

700] that we observed (Figure 3.5) should be reduced when Atg13 is bound to Vac8. To investigate this possibility, we carried out liposome sedimentation assays using recombinant purified

97 Atg13[571-700] and Vac8[10-515] at 50 and 150 mM NaCl. Atg13 was incubated with either

Vac8[10-515] or buffer prior to mixing with liposomes to allow for Atg13[571-700]-Vac8[10-

515] complex formation. Vac8[10-515] did not pellet in the presence of liposomes at any salt concentration tested indicating that this construct does not bind liposomes (Figure 3.10). At 50 mM NaCl, Vac8[10-515] reduced Atg13[571-700] liposome binding by 45% (Figure 3.10A and

Figure 3.10B). At 150 mM NaCl, Vac8[10-515] reduced the amount of Atg13[571-700] in the pellet essentially to background levels, demonstrating that there was no liposome binding by

Atg13[571-700] in the presence of Vac8[10-515] (Figure 3.10C and Figure 3.10D). These data demonstrate that Vac8[10-515] binding to Atg13[571-700] inhibits liposome binding by the latter and suggest that the Vac8 and liposome binding sites on Atg13 are mutually exclusive despite utilizing distinct amino acids.

3.3.7 The Atg13 C-terminal region is necessary for localization of the protein near the PAS and vacuole during starvation

Our finding that the Atg13 region 571-700 alternates binding between phospholipids and the vacuole membrane-associated protein Vac8 suggests that the protein-membrane network, which is organized by the Atg13 hub-like IDR, is maintained in proximity of the vacuole via the Vac8- binding capability of this region. To probe such a role of the 571-700 region in Atg13, we constructed a yeast strain expressing the truncated Atg13[1-570]-GFP with RFP-tagged Ape1 (a marker for the PAS) and compared it with the strain expressing the full-length Atg13-GFP. Similar to previous IDR mutants, Atg13[1-570]-GFP was partially defective in autophagy activity (Figure

3.11A). Furthermore, this construct did not colocalize efficiently with RFP-Ape1 relative to the

98 full-length protein (Figure 3.11B), suggesting that the yeast-specific Atg13 C terminus is needed for the proximity of the protein near the PAS, which is close to the vacuole. To further confirm this result, we analyzed colocalization of Atg13-GFP and Atg13[1-570]-GFP relative to the FM 4-

64-labeled vacuolar membrane. Again, the truncation mutant Atg13[1-570]-GFP exhibited a localization further away from the vacuole than the wild-type Atg13 (Figure 3.11C-E). Together these data show that the Atg13 C terminus is required for perivacuolar localization of the protein during starvation.

99 3.4 Discussion

Numerous cellular processes rely on optimal spatiotemporal organization of biological molecules

(proteins, DNA, RNA, or membranes). Because intricate assembly and efficient interaction of structurally diverse components must be free of steric hindrances, these processes are often mediated by a backbone or a hub that exhibits a high architectural plasticity, which is an embedded feature of intrinsically disordered regions301, 302. Multiple studies in recent years showed that the

IDR from yeast Atg13 functions as such a hub. So far, Atg13 IDR was shown to be essential for assembly of two different Atg17 dimers, the EAT domain of the Atg1 kinase, and the peripheral vacuolar protein Vac8293, 295, 298, 303.

Here we show that the Atg13 IDR also incorporates phospholipid membranes into the protein network assembly. The interaction between phospholipids and the Atg13 IDR is electrostatic, where negatively charged phospholipids bind to specific, positively charged residues located in 2 Phe-containing motifs (640KFK and 683KFHK) in the 571-700 region of Atg13.

Mutagenic alteration that puts glutamic acid in these motifs results in substantially weakened liposome binding along with autophagy-deficient phenotypes, despite normal and even more efficient binding to Vac8 in in vitro and in vivo conditions, respectively (Figure. 3.5-3.10). Such results suggest that the membrane binding capability alone of the 571-700 region in Atg13 is essential for efficient autophagy. At the same time, the interaction of Atg13 with Vac8 is also required for autophagy303. These findings collectively show that the Atg13 C terminus must be able to interact with both binding partners (phospholipids and Vac8) to completely fulfill its function in autophagy.

100 The linear motifs for phospholipid binding in the C-terminal IDR of Atg13 overlap with the Vac8-binding domain. Overlapping binding regions are not unusual in IDRs. A well-known hub, TP53/p53, carries multiple overlapping motifs in its disordered C-terminal sequence304.

Another example is the disordered N-terminal domain of KNL1 ( kinetochore scaffold 1), a protein that binds microtubules at two sites, which overlap with the PPP1 (protein phosphatase 1) biding sites305. A purpose of these overlapping regions is to function as molecular switches, because they exhibit a mutually exclusive interaction.

Indeed, in analogy to TP53 and KNL1, we found that the phospholipid- and Vac8-binding of Atg13 is mutually exclusive (Figure 3.10), suggesting that the Atg13 C terminus reversibly switches between attachment to Vac8 and membrane. It is plausible to assume that the hyperphosphorylated Atg13 IDR binds the Vac8 groove, where cationic residues were found to be essential for the Atg13-Vac8 interaction. This would mask the phospholipid-binding motifs and prevent them from attaching to a membrane. A regulatory mechanism, possibly dephosphorylation of Atg13 upon starvation that removes negatively charged phosphoryl groups, could regulate a release of the positively charged phospholipid-binding motifs (640KFK and 683KFHK) from the

Vac8 groove, and thereby facilitate their attachment to a negatively charged membrane (Figure

3.12).

Elucidation of a regulatory switching mechanism is out of the scope of this study and remains to be solved by future research. Finding a purpose of the phospholipid-binding capacity of the

Atg13 C terminus near the vacuole is another task for future studies. At this point, we can only speculate that this newly discovered function of Atg13 might navigate growth of the phagophore

101 in a direction towards the vacuole, which would later facilitate efficient autophagosome-vacuole fusion.

3.5 Materials and Methods

3.5.1 Overexpression, purification, and characterization of recombinant Atg13[571-700] and mutants

S. cerevisiae Atg13[571-700] was subcloned into pHis2 and the pET His6 tobacco etch virus ligation-independent cloning vector (1B) (Addgene, 29653; deposited by Scott Gradia ).

Atg13[647-700], Atg13[571-682], and Atg13[571-640] were also subcloned into this same vector. All mutants were generated from Atg13[571-700] pHis2 using Q5 mutagenesis

(NEB). Atg13 constructs were transformed into E. coli BL21 (DE3) RIL cells (Invitrogen). Cells were grown in TB medium at 37°C to an OD600 of 1.2, protein expression was induced with 1 mM isopropyl-beta-D-thiogalactopyranoside, and cells were subsequently grown for 18 h at

18°C. Cells were harvested and pellets were stored at -80°C. Cell pellets were thawed and resuspended in 50 mM Tris, pH 7.4, 500 mM NaCl, 0.1% Triton X-100 containing an EDTA-free mini protease inhibitor tablet (Roche). Cells were lysed with three passes through a French press

(Thermo Electron). Lysates were cleared by centrifugation and the cleared supernatant was added to TALON resin (Clontech) and incubated at 4°C for 1 h. The resin was washed with 50 mM Tris, pH 7.4, 500 mM NaCl, and the protein was eluted using 50 mM Tris, pH 7.4, 500 mM NaCl, 200 mM imidazole. Elutions containing protein were pooled and further purified using a HiLoad

102 16/600 Superdex 75-pg column (GE Healthcare) equilibrated in 20 mM Tris, pH 7.4, 100 mM

NaCl, 0.2 mM Tris(2-carboxyethyl)phosphine (TCEP).

3.5.2 Liposome sedimentation assay

Folch fraction type I lipids isolated from bovine brain (Sigma-Aldrich) and synthetic liposomes made using L-α-phosphatidylcholine (Avanti), 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-L- serine (Avanti), L-α-phosphatidylethanolamine (Avanti), L-α-phosphatidylinositol (Avanti), 1,2- dioleoyl-sn-glycero-3-phospho-(1'-myo-inositol-3'-phosphate) (Avanti), and cholesterol (Avanti) were dried under a nitrogen stream for 30 min and then for 18 h in a vacuum oven. Dried Folch was resuspended in 20 mM Tris, pH 7.4, 200 mM NaCl, 0.2 mM TCEP to a final concentration of 0.5 mg/mL. Synthetic liposomes were resuspended in 20 mM Tris, pH 7.4,

50 mM NaCl, 0.2 mM TCEP to a final concentration of 1.9 mg/mL. Lipids were subjected to freeze-thaw by incubating at -80°C for 10 min and then thawing in a water bath. Liposomes were extruded using an Avanti Mini Extruder with the appropriate size membrane. Purified Atg13[571-

700] (25 μL of 5 μM) was mixed with 25 μL of 0.5 mg/mL folch liposomes or 1.9 mg/mL synthetic liposomes. For Atg13[571-700] liposome binding in the presence of Vac8[10-515], 12.5

µL of 24 µM Atg13 was mixed with either 12.5 µL of 24 µM Vac8[10-515] or buffer and incubated for 30 min on ice to allow for complex formation. Folch liposomes (25 µL) were then added at a final concentration of 0.25 mg/ml liposomes and 6 µM protein. Liposome and protein mixture were incubated at 4°C for 1 h, followed by centrifugation at 45,000 rpm for 40 min at 4°C using a

TLA45 rotor. Supernatants were removed, and an equal volume of buffer was added to resuspend

103 the pellets. Samples were subjected to SDS-PAGE for analysis and bands were quantified using

Image Lab v 5.1 (BioRad).

3.5.3 Yeast in vivo assays

Saccharomyces cerevisiae strains WLY176 and JMY347 were used to generate atg13∆, ATG13-PA, full length ATG13-GFP and truncated ATG13[1-570]-GFP strains as previously described275, 276 (Table 3.1).

Genomic point mutations ATG13-PA F641G I645G, F641E I645E and K683E H685E K686E were generated as previously described277 (Table 3.1). Yeasts cultures were grown in YPD medium (1% [w:v] yeast extract, 2% [w:v] peptone, and 2% [w:v] glucose) to mid-log phase and then samples were collected.

Strains grown in YPD were shifted to minus nitrogen medium (0.17% yeast nitrogen base without ammonium sulfate or amino acids, and 2% [w:v] glucose) for the indicated time points and then collected.

Pho8Δ60 and western blot analyses were performed as previously described254, 306. Western blot densitometry quantification was performed using ImageJ software.

3.5.4 Fluorescence microscopy

Yeast cells were grown to OD600 ~0.5 in YPD and shifted to minus nitrogen medium for autophagy induction. For vacuolar staining, cells were grown in YPD with FM 4-64 (30 μM) for 30 min and then washed to remove excess dye. Images were collected on a DeltaVision Elite deconvolution microscope (GE Healthcare/Applied Precision) with a 100x objective and a CCD camera

(CoolSnap HQ; Photometrics). For quantification of full length Atg13-GFP or truncated Atg13[1-

570]-GFP colocalization with either RFP-Ape1 or FM 4-64-dyed vacuolar membrane, stacks of

20 image planes were collected with a spacing of 0.2 mm to cover the entire yeast cell. Analysis was performed on an average projection of the imaging planes on ImageJ software.

104 3.5.5 His-tag affinity isolation experiments

Cells (60 OD600 units) starved for 1 h in SD-N medium were lysed in 0.7 ml of lysis buffer (1x

PBS [137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM K2HPO4, pH 7.4], 0.2 M sorbitol, 1 mM MgCl, 0.5% Triton X-100, 1 mM PMSF, Complete EDTA-free protease inhibitor [Roche]) with glass beads in 6 cycles of vortexing for 45 s with 2-min interval of incubation on ice between each vortexing cycle. Cell debris were removed by centrifugation at 4000 x g for 5 min. An aliquot of the supernatant fraction (20%) was TCA precipitated as the Input. The remaining supernatant was incubated with 100 µL Ni-NTA agarose for 3 h at 4°C. After three washes with ice-cold lysis buffer, the proteins were eluted by incubating the agarose for 15 min at 55°C with SDS-PAGE buffer containing 150 mM imidazole. The eluted proteins were analyzed by western blot with anti-polyhistidine monoclonal and anti-Vac8 polyclonal antibodies.

3.5.6 Isothermal titration calorimetry

ITC was performed using a VP-ITC (Microcal). All proteins were prepared in 25 mM Tris, pH

7.5, 150 mM NaCl, 5 mM beta-mercaptoethanol for ITC experiments. Recombinant purified

Atg13[571-700] wild-type and mutants at approximately 50 µM were titrated into Vac8[10-515] at 5.1 µM at 24°C. Data were analyzed using Origin (Microcal).

3.5.7 Statistics and reproducibility

Analysis and image processing of microscopy data were carried out using softWoRx software (GE

Healthcare). Sample sizing for cellular imaging was chosen to be the minimum number of

105 independent experiments required for statistically significant results. Western blot images were quantified using ImageJ software. Statistical analyses were performed using GraphPad Prism 6.

3.6 Acknowledgements

This work was supported by NIH grant GM053396 (DJK), GM113132 (MJR) and GM128663 (MJR).

106 Table 3.1. S. cerevisiae strains used in this study. Name Genotype Reference

WLY176 SEY6210 pho13∆ pho8::pho8∆60 253

JMY347 WLY176 pZEO1-pho8∆60 pCu-GFP-ATG8::LEU2 307

DGY056 WLY176 ATG13-PA::TRP1 This study

DGY057 WLY176 ATG13-PA F641G I645G::TRP1 This study

DGY058 WLY176 ATG13-PA F641E I645E::TRP1 This study

DGY059 WLY176 ATG13-PA K683E H685E K686E::TRP1 This study

DGY060 WLY176 atg13Δ::HIS3 This study

DGY061 JMY347 ATG13-PA::TRP1 This study

DGY062 JMY347 ATG13-PA F641G I645G::TRP1 This study

DGY063 JMY347 ATG13-PA F641E I645E::TRP1 This study

DGY064 JMY347 ATG13-PA K683E H685E K686E::TRP1 This study

DGY065 JMY347 atg13Δ::HIS3 This study

DGY066 WLY176 ATG13-GFP::HIS3 This study

DGY067 WLY176 ATG13-GFP [1-570]::HIS3 This study

DGY068 WLY176 ATG13-GFP::HIS3 RFP-APE1::TRP1 This study

DGY069 WLY176 ATG13-GFP [1-570]::HIS3 RFP-APE1::TRP1 This study

YCY131 SEY6210 atg1∆, atg2∆, atg3∆, atg4∆, atg5∆, atg6∆, atg7∆, 300 atg8∆, atg9∆, atg10∆, atg11∆, atg12∆, atg13∆, atg14∆, atg16∆, atg17∆, atg18∆, atg19∆, atg20∆, atg21∆, atg23∆, atg24∆, atg27∆, atg29∆, atg31∆::BLE MKO YCY131 pVac8 pCu6xHis-scrambled sequence Empty::TRP1 This study pVac8

107 pCu6xHis- SS Empty

MKO YCY131 pVac8 pCu6xHis-ATG13[571-700] WT scrambled This study pVac8 sequence::TRP1 pCu6xHis- WT SS MKO YCY131 pVac8 pCu6xHis-ATG13[571-700] F641G I645G This study pVac8 scrambled sequence::TRP1 pCu6xHis- FIGG SS MKO YCY131 pVac8 pCu6xHis-ATG13[571-700] F641E I645E This study pVac8 scrambled sequence::TRP1 pCu6xHis- FIEE SS MKO YCY131 pVac8 pCu6xHis-ATG13[571-700] K683E H685E This study pVac8 K686E scrambled sequence::TRP1 pCu6xHis- KEHEKE SS MKO YCY131 pVac8 pCu6xHis-ATG13[571-700] D651R Q666R This study pVac8 P667R scrambled sequence::TRP1 pCu6xHis- DQPRRR SS MKO YCY131 pCu6xHis-ATG13[571-700] scrambled This study pCu6xHis- sequence::TRP1 WT SS

108 Table 3.2. Dissociation constants for the Atg13[571-700] wild-type and mutant proteins titrated into Vac8[10-515] as determined by ITC.

109

Figure 3.1 1D 1H NMR spectrum of Atg13[571-700]. The lack of dispersion in the amide region of the spectrum is indicative that this region is intrinsically disordered.

Figure 3.2 Circular dichroism. Circular dichroism spectroscopy on Atg13[571-700] (blue), Folch liposomes (green) and Atg13[571-700] with liposomes (red) in 200 mM NaCl. No increase in CD signal was observed

110 at 222 nm indicating that Atg13[571-700] does not gain helical content upon binding to liposomes.

Figure 3.3 Atg13[571-700] binds negatively charged membranes. A. Liposome sedimentation assay of Atg13[571-700] with Folch liposomes containing diameters from 1.0 µm to 0.4 µm. B. Densitrometric quantification of panel (A). C. Liposome sedimentation assay of Atg13[571-700] with 0.6-µm Folch liposomes at varying concentrations of NaCl. D. Densitrometric quantification of panel (C). E. Liposome sedimentation assay of Atg13[571-700] with liposomes composed of PC, PS, PtdIns (PI) and PtdIns3P (PI3P). F. Liposome sedimentation

111 assay of Atg13[571-700] with liposomes composed of PC, PS, PE, Chol, PtdIns and PtdIns3P. g, Densitrometric quantification of panel (F). Error bars represent the SD from three experiments. ANOVA, *P<0.05, ***P<0.001.

112

Figure 3.4 Autophagy activity of Atg13 mutants. A. Amino acid sequence of Atg13[571-700] created in the BioEdit Sequence Alignment Editor308. The phospholipid-binding motifs identified in this study are highlighted by gray rectangles. The Atg13 sequence forming the interface with Vac8 is highlighted by a purple rectangle. Asterisks mark Ser residues that are phosphorylated by Tor in nutrient-rich conditions. B. Autophagy activity was determined using the GFP-Atg8 processing assay in wild-type (WT), Atg13F641,I645E, Atg13K683,H685,K686E and Atg13F641,I645,K683,H685,K686E mutants under nutrient-rich conditions and after 1 h of nitrogen starvation. Error bars indicate the standard deviation of 3 independent experiments. ANOVA, *P <0.05, **P <0.01, n.s. no significant difference. C. Autophagy activity was measured with the prApe1 processing assay in vac8∆ deletion strains expressing either WT Atg13, Atg13F641,I645E, or Atg13F641,I645G mutants or deleted for ATG13 under nutrient-rich

113 conditions and after 0.5 and 1 h of nitrogen starvation. Error bars indicate the standard deviation of 3 independent experiments. ANOVA, ***P <0.001. D. Autophagy activity was measured with the prApe1 processing assay in the vac8∆ deletion strain expressing either WT Atg13, or the Atg13K683,H685,K686E mutant or deleted for ATG13 under nutrient-rich conditions and after 0.5 and 1 h of nitrogen starvation. Error bars indicate the standard deviation of 3 independent experiments. Student’s t-test, n.s no significant difference.

114

Figure 3.5 Atg13 utilizes two distinct regions for liposome binding. A. Liposome sedimentation assay of recombinant Atg13[571-700], [571-682], [571-640], and [647-700] in the absence of lipid or with 0.6-µm Folch liposomes at varying concentrations of NaCl (top) and densitrometric quantification (bottom). B. Liposome sedimentation assay of recombinant Atg13 mutants F641E, I645E; K640A, K642A; F641E, I645E, K640A, K642A;

115 and K683E, H685E, K686E in the absence of lipid or with 0.6-µm Folch liposomes at varying concentrations of NaCl (top) and densitrometric quantification (bottom). Error bars represent the SD from three individual trials. ANOVA, *** P<0.001.

116

Figure 3.6 Schematic illustration of Atg13 mutants and summary of liposome binding assays.

117

Figure 3.7 Atg13 IDR mutants reduce autophagy activity. A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT), atg13∆ and F641E I645E, F641G I645G and K683E H685E K686E mutants under nutrient-rich conditions (+N) and after 3 h of nitrogen starvation (-N 3 h). Error bars indicate the standard deviation of 3 independent

118 experiments. Student’s t-test, *P<0.05 ***P <0.001. B. Autophagy was determined by the GFP- Atg8 processing assay in WT, atg13∆ and F641E I645E, F641G I645G and K683E H685E K686E mutants under nutrient-rich conditions and after 1 h of nitrogen starvation. A representative image is shown. Error bars indicate the standard deviation of 3 independent experiments. ANOVA, ***P <0.001.

Figure 3.8 Atg13[571-700] interacts with Vac8. A. B. His-tag affinity-isolation experiments. Top panels show schematic representation of the different Atg13 constructs that were immobilized on the Ni-NTA agarose to detect their interaction with Vac8 after affinity isolation (AI). MKO cells transformed with the pVac8(426) plasmid and

119 pCu6xHis-Ss(424), pCu6xHis-Atg13[571-700](424) or pCu6xHis-Atg13[571-700]-Ss(424) plasmid were cultured in nutrient-rich conditions to OD600 ~1 and then shifted to nitrogen- starvation medium for 1 h. The wild-type and mutated variants of proteins were separated by SDS- PAGE and detected with anti-Vac8 or anti-His antibodies (lower panels). C. Atg13[571-700] (green and purple) modeled on the Nvj1-Vac8 crystal structure (PDB ID: 5XJG). Positively (blue) and negatively (red) charged residues in the Vac8 (gray) homodimer are highlighted. The position of K640, K642, and Y670 (the latter homologous to Y321 in Nvj1 is denoted based on the Nvj1- Vac8 crystal structure. The position of K683, H685, and K686 in the Atg13 IDR is approximate. D. Raw ITC data (top) and binding isotherms (bottom) for Atg13[571-700] WT, F641E I645E and K683E H685E K686E titrated into Vac8[10-515].

120

Figure 3.9 Interaction between Atg13 and Vac8. A. Schematic representation of the Atg13 constructs that were used in the experiments with whole cell lysates. B-D. The plasmid pCu6xHis-Atg13[571-700](424) or pCu6xHis-Atg13[571-700]- Ss(424) was transformed into the MKO strain with or without the pVac8(426) plasmid

121 overexpressing Vac8. MKO cells were cultured in nutrient-rich conditions to OD600 ~1 and then shifted to nitrogen starvation medium for 1 h. The lysates were TCA precipitated, and the proteins were separated by SDS-PAGE and detected with the indicated antibody or antisera.

122

Figure 3.10 Vac8 inhibits Atg13[571-700] liposome binding. A, B. Atg13[571-700], Vac8[10-515] and a mixture of both proteins were incubated with Folch liposomes in buffer containing 50 mM NaCl and subjected to centrifugation. Pellet and supernatant fractions were subjected to centrifugation. The percent of Atg13[571-700] in the pellet was quantified using densitometry. C, D. Same as a,b except performed in the presence of 150 mM

123 NaCl. Error bars indicate the standard deviation of 4 independent experiments. ANOVA, ***P <0.001.

124

Figure 3.11 Autophagy activity and cellular localization of the truncated Atg13 [1-570] A. Autophagy activity was measured by the Pho8Δ60 assay in wild-type (WT), Atg13-GFP full length, Atg13-GFP[1-570] and atg13∆ cells under nutrient-rich conditions (+N) and after 3 h of nitrogen starvation (-N). Error bars indicate the standard deviation of 3 independent experiments. ANOVA, *P<0.05, **P <0.01, ***P <0.001. B. RFP-Ape1 and Atg13-GFP full length or Atg13- GFP[1-570] puncta colocalization was measured through fluorescence microscopy after 1 h of nitrogen starvation (-N 1 h). Representative image planes are shown. C. The ratio between RFP- Ape1 and Atg13-GFP full length or Atg13-GFP[1-570] colocalizing puncta in cells expressing both RFP-Ape1 and Atg13-GFP. Error bars indicate the standard deviation of 3 independent

125 experiments. Students t-test. ***P <0.001. D. The vacuole membrane dye FM 4-64 and Atg13- GFP full length or Atg13-GFP[1-570] puncta colocalization was measured through fluorescence microscopy after 1 h of nitrogen starvation (-N 1 h). Representative image planes are shown. E. The ratio between Atg13-GFP full length or Atg13-GFP[1-570] puncta co-localizing with the FM 4-64-stained vacuolar membrane is presented. Error bars indicate the standard deviation of 3 independent experiments. Students t-test. ***P <0.001.

126

Figure 3.12 Model of proposed mechanism for a reversible switch in binding of the Atg13 C terminus from Vac8 to phospholipid membranes. The phosphorylated 571-700 region of Atg13 from S. cerevisiae binds to the Vac8 groove where cationic residues are critical for interaction. This masks lipid-binding motifs and prevents them from attaching to the membrane. A regulatory mechanism, perhaps dephosphorylation upon starvation that removes negatively charged phosphoryl groups, could facilitate a release of the Atg13 positively charged motifs (640KFK and 683KFHK) from Vac8, and allow their attachment to a negatively charged phospholipid membrane.

127 Chapter 4 Summary4

Autophagy is a conserved catabolic recycling pathway involving the degradation of multiple cytoplasmic components during standard physiological conditions and in response to different types of stress, such as nutrient starvation. Macroautophagy/autophagy involves the sequestration of cytoplasm within double-membrane vesicles termed autophagosomes. Upon maturation, autophagosomes fuse with the vacuole or lysosomes, leading to degradation of their cargo by resident hydrolases. Cargo degradation produces macromolecules such as amino acids, which are subsequently recycled back into the cytoplasm for reuse. Autophagy dysregulation has been involved in the development of a diverse range of diseases such as cancer and different neurodegenerative disorders. Several ATG genes and their corresponding proteins have been characterized based on their function(s) in regulating the various stages of autophagy. Upon nutrient starvation, ATG gene transcription is dramatically upregulated to increase Atg protein synthesis and autophagy activity. While several breakthroughs have been made in understanding how autophagy is regulated, we are still learning the specific mechanisms behind autophagy regulation at the post-transcriptional and structural level.

In this chapter, I summarize findings of chapters 2 and 3, and discuss future directions.

4 This chapter includes a modified version of the article Damian Gatica, Guowu Hu, Xu Liu, Nannan Zhang, Peter R. Williamson and Daniel J. Klionsky. The Pat1-Lsm complex prevents 3' to 5' degradation of a specific subset of ATG mRNAs during nitrogen starvation-induced autophagy. (2019) Autophagy (doi: 10.1080/15548627.2019.1587262).

128 4.1 The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen Starvation-Induced

Autophagy

In chapter 2 we characterized a new post-transcriptional regulation pathway for several ATG mRNA that is necessary for robust autophagy induction during nitrogen starvation-induced autophagy. We first described a decrease in autophagy activity in PAT1 and LSM1 deletion strains after nitrogen starvation, which correlates with decreased protein and ATG mRNA levels of the essential ATG genes ATG1, ATG2, ATG7 and ATG9. Preventing efficient exosome-mediated 3' to

5' mRNA degradation by deleting SKI3, restores autophagy activity in the PAT1 deletion strain, as well as ATG1, ATG2, ATG7 and ATG9 protein and mRNA levels to those observed in a wild-type strain. Measuring the mRNA levels of all known yeast ATG genes in a pat1Δ strain, revealed that the Pat1-Lsm complex affects only a subset of ATG mRNAs during nitrogen starvation; the levels of several other ATG mRNA transcripts, such as ATG17 and ATG18, remain unaffected by PAT1 deletion.

Because Pat1 is rapidly dephosphorylated at serine residues S456 and S457 under conditions of glucose starvation, we wondered if Pat1 was also dephosphorylated during nitrogen starvation, and if this dephosphorylation could be relevant to autophagy regulation. Indeed, Pat1 is dephosphorylated at the S456 and S457 residues during nitrogen starvation. Furthermore, a phosphomimetic Pat1 mutant (Pat1S456,457E) displays a similar decrease in autophagy activity, as well as ATG1, ATG2, ATG7 and ATG9 protein and mRNA levels as that seen in a PAT1 deletion strain. Conversely, when ATG17 and ATG18 mRNA levels are measured in the Pat1S456,457E strain, they fail to show any difference compared to a strain expressing wild-type Pat1. This difference between ATG mRNAs appears to be due to the ability of Pat1 to bind specific transcripts, as observed when binding is measured through mRNA immunoprecipitation as well as a protein-

129 RNA bimolecular fluorescence complementation assay. Once again, preventing efficient 3' to 5' mRNA degradation by deleting SKI3 restores the autophagy activity, as well as ATG1, ATG2,

ATG7 and ATG9 protein and mRNA levels in the Pat1S456E,S457E strain to those observed in a wild- type strain.

Because the Pat1-Lsm complex binds preferentially to the 3' UTR of mRNA, we decided to switch the 3' UTR of ATG1 mRNA, which shows a decrease when PAT1 is deleted, with the 3'

UTR of ATG17 and ATG18 mRNA, which remain unaffected by PAT1 deletion. Switching the

ATG1 mRNA 3' UTR with that of either gene prevents the PAT1 deletion-mediated decrease of

ATG1 mRNA and protein levels after nitrogen starvation. Finally, we determined if the Pat1-Lsm complex effect on ATG mRNA was conserved in humans. The siRNA-mediated knockdown of the PAT1 ortholog PATL1 leads to a decrease in ATG2 and ATG9A mRNA levels. Consistent with our yeast data, a PATL1 phosphomimetic mutant shows decreased ATG2 and ATG9A mRNA levels under autophagy-inducing conditions, whereas a PATL1 nonphosphorylatable mutant displays increased autophagy activity.

Our study further elucidates how autophagy is regulated at the post-transcriptional level by different mRNA binding proteins. However, important questions remain about which other mRNA-binding protein are involved in autophagy regulation, as well as understanding the specific mechanisms by which these mRNA-binding proteins recognize their target mRNAs.

130 4.2 The carboxy terminus of yeast Atg13 binds phospholipid membrane via motifs that

overlap with the Vac8-interacting domain

In chapter 3 we identified two sets of phospholipid-binding residues in the IDR of Atg13, a key member of the autophagy initiation complex. These phospholipid-binding residues overlap with the Vac8-binding domain of Atg13 and are required for efficient autophagy activity during nitrogen starvation.

The Atg13 C terminus is predicted to be an intrinsically disordered region harboring a binding site for the vacuolar membrane protein Vac8. Recent reports suggest Atg13 acts as a hub to assemble the initiation complex, as well as participating in membrane recognition288, 293, 294. To test whether part of the Atg13 C-terminal IDR interacts with phospholipids, Atg13[571-700] was purified and incubated with Folch-derived unilamellar vesicles for liposome sedimentation assays.

Atg13[571-700] was able to bind liposomes through electrostatic interactions because increasing concentrations of NaCl disrupted the interaction. Incubating Atg13[571-700] with liposomes containing different concentrations of polar phospholipids revealed that Atg13[571-700] binds preferentially negatively charged phospholipids.

Truncating Atg13[571-700] in different regions containing positively charged residues indicated that the Atg13[641-700] region was important for liposome binding. This region contains two sets of positively charged residues, 683KFHK and 640KFKSSI, that when mutated to the negatively charged residue glutamate, decreased liposome binding.

We then measured autophagy activity of the Atg13 F641E I645E and K683E H685E

K686E mutants through the Pho8Δ60 and GFP-Atg8 processing assay. Both set of mutations resulted in a significant decrease in autophagy for both mutations when compared to wild-type

Atg13.

131 Because the phospholipid-binding motifs identified in our study were within the Atg13

Vac8-binding region we decided to investigate the effect these mutations could have in Vac8 binding through affinity-isolation experiments. Both Atg13 F641E I645E and K683E H685E

K686E showed increased Vac8 binding compared to wild-type Atg13[571-700], which could be explained by the cationic residues in the Vac8 groove that are responsible for Atg13 interaction.

Testing the binding affinity of Atg13 mutants with Vac8 using isothermal calorimetry revealed that the dissociation constants for wild-type and mutant Atg13[571-700] were very similar, suggesting that whereas the Atg13 residues required for liposome binding are not required for

Vac8 binding, Atg13 binding to membranes or Vac8 might be mutually exclusive because both regions are overlapping. To test this possibility Atg13[571-700] liposome sedimentation assays were carried out in the presence of Vac8[10-515], which is unable to bind liposomes. As expected,

Atg13[571-700] failed to bind liposomes efficiently in the presence of Vac8[10-515], indicating

Vac8 and liposome binding sites on Atg13 are mutually exclusive despite utilizing distinct amino acids.

The latter result led us to propose that Atg13 maintains vacuolar proximity by interacting with the vacuole membrane-associated protein Vac8. To test this hypothesis, we used microscopy to check co-localization between wild-type Atg13-GFP or truncated Atg13[1-570]-GFP and either

RFP-Ape1 (a marker for the PAS) or FM 4-64-labeled vacuolar membrane. Our results showed that whereas wild-type Atg13-GFP co-localized with RFP-Ape1 and the vacuole, Atg13[1-570]-

GFP failed to do so, suggesting that the Atg13 C terminus is required for its perivacuolar localization during starvation.

132

4.3 Discussion and perspectives

Recent years have highlighted the importance of understanding the mechanisms behind autophagy regulation at every level, from the transcriptional regulation of the ATG genes to the molecular interactions between the different Atg proteins. Both studies presented in this dissertation further our understanding of how autophagy is regulated, first, at the post-transcriptional level and second, at the molecular and structural level. However, we are still only beginning to understand the specific roles different mRNA-binding proteins have in regulating autophagy by preventing ATG mRNA degradation or inducing efficient ATG transcript translation during autophagy-inducing conditions. Our results highlight the importance of the Pat1-Lsm mRNA-binding complex in preventing the degradation of a specific subset of ATG mRNAs. However, we still do not know the precise mechanism by which some of these ATG mRNAs are recognized, while others remain unaffected. Point mutations in the 3' UTR of some of these ATG transcripts, along with RNA immunoprecipitation experiments could potentially uncover more information regarding the specificity of the Pat1-Lsm complex-binding targets. Interestingly, our results also suggest that some ATG mRNAs, such as ATG3 and ATG8, are differentially regulated by the Pat1-Lsm complex. In the case of ATG8, this is the only ATG mRNA whose transcript levels are significantly increased during nutrient-rich conditions in the absence of Pat1. The latter combined with the central role of Atg8 in regulating the autophagic process, emphasizes the need for further experiments to understand this interaction. A continued analysis of other mRNA-binding proteins, that are also Pat1 binding partners, such as Dhh1, could further our understanding of how these

ATG mRNAs are differentially regulated during nutrient-rich and nitrogen starvation conditions.

133 Similarly, there is a need to understand not only how the different Atg proteins interact with one another but to comprehend the hierarchy of each protein in each individual step of autophagy. Our data indicating that Atg13 competes for binding between the Vac8 vacuolar membrane protein and negatively charged membranes, opens interesting questions regarding which event takes place first during the initiation of the autophagic process. We can speculate that

Vac8 binding would occur first, in order to localize Atg13 close to the vacuolar membrane. After proper localization Atg13 would act as a scaffold for recruiting other Atg proteins involved in the initiation complex. Later, as the phagophore increases size through Atg9 membrane trafficking, we would expect Atg13 binding to the negatively charged membranes. Binding experiments, such as immunoprecipitation and yeast two-hybrid, between Atg13 and other known interacting partners, in a vac8 deletion background, could highlight the importance of Atg13-binding to Vac8 in regulating the proper localization of the initiation complex and its effects on autophagy activity.

Further research will be necessary to reveal answers to all these questions and to provide additional knowledge that permits us to use this information in preventing and treating human diseases.

134 Bibliography

1. Feng, Y., He, D., Yao, Z. & Klionsky, D.J. The machinery of macroautophagy. Cell Res 24, 24-41 (2014). 2. Gatica, D., Chiong, M., Lavandero, S. & Klionsky, D.J. Molecular mechanisms of autophagy in the cardiovascular system. Circ Res 116, 456-467 (2015). 3. Kaushik, S. & Cuervo, A.M. Chaperone-mediated autophagy: a unique way to enter the lysosome world. Trends Cell Biol 22, 407-417 (2012). 4. Mijaljica, D., Prescott, M. & Devenish, R.J. Microautophagy in mammalian cells: revisiting a 40-year-old conundrum. Autophagy 7, 673-682 (2011). 5. Parzych, K.R. & Klionsky, D.J. An overview of autophagy: morphology, mechanism, and regulation. Antioxid Redox Signal 20, 460-473 (2014). 6. He, C. & Klionsky, D.J. Regulation mechanisms and signaling pathways of autophagy. Annu Rev Genet 43, 67-93 (2009). 7. Jin, M., Liu, X. & Klionsky, D.J. SnapShot: Selective autophagy. Cell 152, 368-368 e362 (2013). 8. Ashrafi, G. & Schwarz, T.L. The pathways of mitophagy for quality control and clearance of mitochondria. Cell Death Differ 20, 31-42 (2013). 9. Hutchins, M.U., Veenhuis, M. & Klionsky, D.J. Peroxisome degradation in Saccharomyces cerevisiae is dependent on machinery of macroautophagy and the Cvt pathway. J Cell Sci 112 ( Pt 22), 4079-4087 (1999). 10. Hung, Y.H., Chen, L.M., Yang, J.Y. & Yang, W.Y. Spatiotemporally controlled induction of autophagy- mediated lysosome turnover. Nat Commun 4, 2111 (2013). 11. Nakatogawa, H. & Mochida, K. Reticulophagy and nucleophagy: New findings and unsolved issues. Autophagy 11, 2377-2378 (2015). 12. Sarkar, S., Ravikumar, B. & Rubinsztein, D.C. Autophagic clearance of aggregate-prone proteins associated with neurodegeneration. Methods Enzymol 453, 83-110 (2009). 13. Kamada, Y. et al. Tor directly controls the Atg1 kinase complex to regulate autophagy. Mol Cell Biol 30, 1049-1058 (2010). 14. Kabeya, Y. et al. Characterization of the Atg17-Atg29-Atg31 complex specifically required for starvation- induced autophagy in Saccharomyces cerevisiae. Biochem Biophys Res Commun 389, 612-615 (2009). 15. Cheong, H., Nair, U., Geng, J. & Klionsky, D.J. The Atg1 kinase complex is involved in the regulation of protein recruitment to initiate sequestering vesicle formation for nonspecific autophagy in Saccharomyces cerevisiae. Mol Biol Cell 19, 668-681 (2008). 16. Kim, J., Huang, W.P., Stromhaug, P.E. & Klionsky, D.J. Convergence of multiple autophagy and cytoplasm to vacuole targeting components to a perivacuolar membrane compartment prior to de novo vesicle formation. J Biol Chem 277, 763-773 (2002). 17. Reggiori, F., Tucker, K.A., Stromhaug, P.E. & Klionsky, D.J. The Atg1-Atg13 complex regulates Atg9 and Atg23 retrieval transport from the pre-autophagosomal structure. Dev Cell 6, 79-90 (2004). 18. Suzuki, K. et al. The pre-autophagosomal structure organized by concerted functions of APG genes is essential for autophagosome formation. EMBO J 20, 5971-5981 (2001). 19. Kamada, Y. et al. Tor-mediated induction of autophagy via an Apg1 protein kinase complex. J Cell Biol 150, 1507-1513 (2000). 20. Yeh, Y.Y., Wrasman, K. & Herman, P.K. Autophosphorylation within the Atg1 activation loop is required for both kinase activity and the induction of autophagy in Saccharomyces cerevisiae. Genetics 185, 871-882 (2010). 21. Kraft, C. et al. Binding of the Atg1/ULK1 kinase to the ubiquitin-like protein Atg8 regulates autophagy. EMBO J 31, 3691-3703 (2012). 22. Kijanska, M. et al. Activation of Atg1 kinase in autophagy by regulated phosphorylation. Autophagy 6, 1168- 1178 (2010).

135 23. Budovskaya, Y.V., Stephan, J.S., Deminoff, S.J. & Herman, P.K. An evolutionary proteomics approach identifies substrates of the cAMP-dependent protein kinase. Proc Natl Acad Sci U S A 102, 13933-13938 (2005). 24. Budovskaya, Y.V., Stephan, J.S., Reggiori, F., Klionsky, D.J. & Herman, P.K. The Ras/cAMP-dependent protein kinase signaling pathway regulates an early step of the autophagy process in Saccharomyces cerevisiae. J Biol Chem 279, 20663-20671 (2004). 25. Stephan, J.S., Yeh, Y.Y., Ramachandran, V., Deminoff, S.J. & Herman, P.K. The Tor and PKA signaling pathways independently target the Atg1/Atg13 protein kinase complex to control autophagy. Proc Natl Acad Sci U S A 106, 17049-17054 (2009). 26. Natarajan, K. et al. Transcriptional profiling shows that Gcn4p is a master regulator of gene expression during amino acid starvation in yeast. Mol Cell Biol 21, 4347-4368 (2001). 27. Talloczy, Z. et al. Regulation of starvation- and virus-induced autophagy by the eIF2alpha kinase signaling pathway. Proc Natl Acad Sci U S A 99, 190-195 (2002). 28. Yan, J. et al. Identification of mouse ULK1, a novel protein kinase structurally related to C. elegans UNC- 51. Biochem Biophys Res Commun 246, 222-227 (1998). 29. Yan, J. et al. Mouse ULK2, a novel member of the UNC-51-like protein kinases: unique features of functional domains. Oncogene 18, 5850-5859 (1999). 30. Hara, T. et al. FIP200, a ULK-interacting protein, is required for autophagosome formation in mammalian cells. J Cell Biol 181, 497-510 (2008). 31. Jung, C.H. et al. ULK-Atg13-FIP200 complexes mediate mTOR signaling to the autophagy machinery. Mol Biol Cell 20, 1992-2003 (2009). 32. Mercer, C.A., Kaliappan, A. & Dennis, P.B. A novel, human Atg13 binding protein, Atg101, interacts with ULK1 and is essential for macroautophagy. Autophagy 5, 649-662 (2009). 33. Ganley, I.G. et al. ULK1.ATG13.FIP200 complex mediates mTOR signaling and is essential for autophagy. J Biol Chem 284, 12297-12305 (2009). 34. Hosokawa, N. et al. Nutrient-dependent mTORC1 association with the ULK1-Atg13-FIP200 complex required for autophagy. Mol Biol Cell 20, 1981-1991 (2009). 35. Chan, E.Y., Longatti, A., McKnight, N.C. & Tooze, S.A. Kinase-inactivated ULK proteins inhibit autophagy via their conserved C-terminal domains using an Atg13-independent mechanism. Mol Cell Biol 29, 157-171 (2009). 36. Kim, J., Kundu, M., Viollet, B. & Guan, K.L. AMPK and mTOR regulate autophagy through direct phosphorylation of Ulk1. Nat Cell Biol 13, 132-141 (2011). 37. Inoki, K., Li, Y., Xu, T. & Guan, K.L. Rheb GTPase is a direct target of TSC2 GAP activity and regulates mTOR signaling. Genes Dev 17, 1829-1834 (2003). 38. Inoki, K., Zhu, T. & Guan, K.L. TSC2 mediates cellular energy response to control cell growth and survival. Cell 115, 577-590 (2003). 39. Loffler, A.S. et al. Ulk1-mediated phosphorylation of AMPK constitutes a negative regulatory feedback loop. Autophagy 7, 696-706 (2011). 40. Alessi, D.R. et al. Characterization of a 3-phosphoinositide-dependent protein kinase which phosphorylates and activates protein kinase Balpha. Curr Biol 7, 261-269 (1997). 41. Sarbassov, D.D., Guertin, D.A., Ali, S.M. & Sabatini, D.M. Phosphorylation and regulation of Akt/PKB by the rictor-mTOR complex. Science 307, 1098-1101 (2005). 42. Manning, B.D., Tee, A.R., Logsdon, M.N., Blenis, J. & Cantley, L.C. Identification of the tuberous sclerosis complex-2 tumor suppressor gene product tuberin as a target of the phosphoinositide 3-kinase/akt pathway. Mol Cell 10, 151-162 (2002). 43. Petiot, A., Ogier-Denis, E., Blommaart, E.F., Meijer, A.J. & Codogno, P. Distinct classes of phosphatidylinositol 3'-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells. J Biol Chem 275, 992-998 (2000). 44. Arico, S. et al. The tumor suppressor PTEN positively regulates macroautophagy by inhibiting the phosphatidylinositol 3-kinase/protein kinase B pathway. J Biol Chem 276, 35243-35246 (2001). 45. Kametaka, S., Okano, T., Ohsumi, M. & Ohsumi, Y. Apg14p and Apg6/Vps30p form a protein complex essential for autophagy in the yeast, Saccharomyces cerevisiae. J Biol Chem 273, 22284-22291 (1998). 46. Kihara, A., Noda, T., Ishihara, N. & Ohsumi, Y. Two distinct Vps34 phosphatidylinositol 3-kinase complexes function in autophagy and carboxypeptidase Y sorting in Saccharomyces cerevisiae. J Cell Biol 152, 519- 530 (2001).

136 47. Obara, K., Sekito, T. & Ohsumi, Y. Assortment of phosphatidylinositol 3-kinase complexes--Atg14p directs association of complex I to the pre-autophagosomal structure in Saccharomyces cerevisiae. Mol Biol Cell 17, 1527-1539 (2006). 48. Furuya, N., Yu, J., Byfield, M., Pattingre, S. & Levine, B. The evolutionarily conserved domain of Beclin 1 is required for Vps34 binding, autophagy and tumor suppressor function. Autophagy 1, 46-52 (2005). 49. Itakura, E., Kishi, C., Inoue, K. & Mizushima, N. Beclin 1 forms two distinct phosphatidylinositol 3-kinase complexes with mammalian Atg14 and UVRAG. Mol Biol Cell 19, 5360-5372 (2008). 50. Matsunaga, K. et al. Two Beclin 1-binding proteins, Atg14L and Rubicon, reciprocally regulate autophagy at different stages. Nat Cell Biol 11, 385-396 (2009). 51. Zhong, Y. et al. Distinct regulation of autophagic activity by Atg14L and Rubicon associated with Beclin 1- phosphatidylinositol-3-kinase complex. Nat Cell Biol 11, 468-476 (2009). 52. Fimia, G.M. et al. Ambra1 regulates autophagy and development of the nervous system. Nature 447, 1121- 1125 (2007). 53. Sun, Q. et al. Identification of Barkor as a mammalian autophagy-specific factor for Beclin 1 and class III phosphatidylinositol 3-kinase. Proc Natl Acad Sci U S A 105, 19211-19216 (2008). 54. Matsunaga, K. et al. Autophagy requires endoplasmic reticulum targeting of the PI3-kinase complex via Atg14L. J Cell Biol 190, 511-521 (2010). 55. Takahashi, Y. et al. Bif-1 interacts with Beclin 1 through UVRAG and regulates autophagy and tumorigenesis. Nat Cell Biol 9, 1142-1151 (2007). 56. Pattingre, S. et al. Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy. Cell 122, 927-939 (2005). 57. Zalckvar, E. et al. DAP-kinase-mediated phosphorylation on the BH3 domain of beclin 1 promotes dissociation of beclin 1 from Bcl-XL and induction of autophagy. EMBO Rep 10, 285-292 (2009). 58. Russell, R.C. et al. ULK1 induces autophagy by phosphorylating Beclin-1 and activating VPS34 lipid kinase. Nat Cell Biol 15, 741-750 (2013). 59. Kim, J. et al. Differential regulation of distinct Vps34 complexes by AMPK in nutrient stress and autophagy. Cell 152, 290-303 (2013). 60. Hailey, D.W. et al. Mitochondria supply membranes for autophagosome biogenesis during starvation. Cell 141, 656-667 (2010). 61. Hayashi-Nishino, M. et al. A subdomain of the endoplasmic reticulum forms a cradle for autophagosome formation. Nat Cell Biol 11, 1433-1437 (2009). 62. Young, A.R. et al. Starvation and ULK1-dependent cycling of mammalian Atg9 between the TGN and endosomes. J Cell Sci 119, 3888-3900 (2006). 63. Yla-Anttila, P., Vihinen, H., Jokitalo, E. & Eskelinen, E.L. 3D tomography reveals connections between the phagophore and endoplasmic reticulum. Autophagy 5, 1180-1185 (2009). 64. Axe, E.L. et al. Autophagosome formation from membrane compartments enriched in phosphatidylinositol 3-phosphate and dynamically connected to the endoplasmic reticulum. J Cell Biol 182, 685-701 (2008). 65. Polson, H.E. et al. Mammalian Atg18 (WIPI2) localizes to omegasome-anchored phagophores and positively regulates LC3 lipidation. Autophagy 6, 506-522 (2010). 66. Karanasios, E. et al. Dynamic association of the ULK1 complex with omegasomes during autophagy induction. J Cell Sci 126, 5224-5238 (2013). 67. Tooze, S.A. & Yoshimori, T. The origin of the autophagosomal membrane. Nat Cell Biol 12, 831-835 (2010). 68. Hamasaki, M. et al. Autophagosomes form at ER-mitochondria contact sites. Nature 495, 389-393 (2013). 69. He, C., Baba, M., Cao, Y. & Klionsky, D.J. Self-interaction is critical for Atg9 transport and function at the phagophore assembly site during autophagy. Mol Biol Cell 19, 5506-5516 (2008). 70. Noda, T. et al. Apg9p/Cvt7p is an integral membrane protein required for transport vesicle formation in the Cvt and autophagy pathways. J Cell Biol 148, 465-480 (2000). 71. Mari, M. et al. An Atg9-containing compartment that functions in the early steps of autophagosome biogenesis. J Cell Biol 190, 1005-1022 (2010). 72. Yamamoto, H. et al. Atg9 vesicles are an important membrane source during early steps of autophagosome formation. J Cell Biol 198, 219-233 (2012). 73. Reggiori, F., Shintani, T., Nair, U. & Klionsky, D.J. Atg9 cycles between mitochondria and the pre- autophagosomal structure in yeasts. Autophagy 1, 101-109 (2005). 74. He, C. et al. Recruitment of Atg9 to the preautophagosomal structure by Atg11 is essential for selective autophagy in budding yeast. J Cell Biol 175, 925-935 (2006).

137 75. Yen, W.L., Legakis, J.E., Nair, U. & Klionsky, D.J. Atg27 is required for autophagy-dependent cycling of Atg9. Mol Biol Cell 18, 581-593 (2007). 76. Orsi, A. et al. Dynamic and transient interactions of Atg9 with autophagosomes, but not membrane integration, are required for autophagy. Mol Biol Cell 23, 1860-1873 (2012). 77. Webber, J.L. & Tooze, S.A. Coordinated regulation of autophagy by p38alpha MAPK through mAtg9 and p38IP. EMBO J 29, 27-40 (2010). 78. Kim, J., Dalton, V.M., Eggerton, K.P., Scott, S.V. & Klionsky, D.J. Apg7p/Cvt2p is required for the cytoplasm-to-vacuole targeting, macroautophagy, and peroxisome degradation pathways. Mol Biol Cell 10, 1337-1351 (1999). 79. Tanida, I. et al. Apg7p/Cvt2p: A novel protein-activating enzyme essential for autophagy. Mol Biol Cell 10, 1367-1379 (1999). 80. Shintani, T. et al. Apg10p, a novel protein-conjugating enzyme essential for autophagy in yeast. EMBO J 18, 5234-5241 (1999). 81. Mizushima, N. et al. A protein conjugation system essential for autophagy. Nature 395, 395-398 (1998). 82. Mizushima, N., Sugita, H., Yoshimori, T. & Ohsumi, Y. A new protein conjugation system in human. The counterpart of the yeast Apg12p conjugation system essential for autophagy. J Biol Chem 273, 33889-33892 (1998). 83. Kuma, A., Mizushima, N., Ishihara, N. & Ohsumi, Y. Formation of the approximately 350-kDa Apg12- Apg5.Apg16 multimeric complex, mediated by Apg16 oligomerization, is essential for autophagy in yeast. J Biol Chem 277, 18619-18625 (2002). 84. Mizushima, N. et al. Mouse Apg16L, a novel WD-repeat protein, targets to the autophagic isolation membrane with the Apg12-Apg5 conjugate. J Cell Sci 116, 1679-1688 (2003). 85. Mizushima, N., Noda, T. & Ohsumi, Y. Apg16p is required for the function of the Apg12p-Apg5p conjugate in the yeast autophagy pathway. EMBO J 18, 3888-3896 (1999). 86. Geng, J. & Klionsky, D.J. The Atg8 and Atg12 ubiquitin-like conjugation systems in macroautophagy. 'Protein modifications: beyond the usual suspects' review series. EMBO Rep 9, 859-864 (2008). 87. Ohsumi, Y. Molecular dissection of autophagy: two ubiquitin-like systems. Nat Rev Mol Cell Biol 2, 211- 216 (2001). 88. Kirisako, T. et al. The reversible modification regulates the membrane-binding state of Apg8/Aut7 essential for autophagy and the cytoplasm to vacuole targeting pathway. J Cell Biol 151, 263-276 (2000). 89. Tanida, I., Ueno, T. & Kominami, E. Human light chain 3/MAP1LC3B is cleaved at its carboxyl-terminal Met121 to expose Gly120 for lipidation and targeting to autophagosomal membranes. J Biol Chem 279, 47704-47710 (2004). 90. Ichimura, Y. et al. A ubiquitin-like system mediates protein lipidation. Nature 408, 488-492 (2000). 91. Fujita, N. et al. The Atg16L complex specifies the site of LC3 lipidation for membrane biogenesis in autophagy. Mol Biol Cell 19, 2092-2100 (2008). 92. Hanada, T. et al. The Atg12-Atg5 conjugate has a novel E3-like activity for protein lipidation in autophagy. J Biol Chem 282, 37298-37302 (2007). 93. Noda, N.N., Fujioka, Y., Hanada, T., Ohsumi, Y. & Inagaki, F. Structure of the Atg12-Atg5 conjugate reveals a platform for stimulating Atg8-PE conjugation. EMBO Rep 14, 206-211 (2013). 94. Romanov, J. et al. Mechanism and functions of membrane binding by the Atg5-Atg12/Atg16 complex during autophagosome formation. EMBO J 31, 4304-4317 (2012). 95. Kabeya, Y. et al. LC3, a mammalian homologue of yeast Apg8p, is localized in autophagosome membranes after processing. EMBO J 19, 5720-5728 (2000). 96. Klionsky, D.J. et al. Guidelines for the use and interpretation of assays for monitoring autophagy (3rd edition). Autophagy 12, 1-222 (2016). 97. Kabeya, Y. et al. LC3, GABARAP and GATE16 localize to autophagosomal membrane depending on form- II formation. J Cell Sci 117, 2805-2812 (2004). 98. Tanida, I., Komatsu, M., Ueno, T. & Kominami, E. GATE-16 and GABARAP are authentic modifiers mediated by Apg7 and Apg3. Biochem Biophys Res Commun 300, 637-644 (2003). 99. Tanida, I., Tanida-Miyake, E., Komatsu, M., Ueno, T. & Kominami, E. Human Apg3p/Aut1p homologue is an authentic E2 enzyme for multiple substrates, GATE-16, GABARAP, and MAP-LC3, and facilitates the conjugation of hApg12p to hApg5p. J Biol Chem 277, 13739-13744 (2002). 100. Weidberg, H. et al. LC3 and GATE-16/GABARAP subfamilies are both essential yet act differently in autophagosome biogenesis. EMBO J 29, 1792-1802 (2010). 101. Klionsky, D.J. The molecular machinery of autophagy: unanswered questions. J Cell Sci 118, 7-18 (2005).

138 102. Darsow, T., Rieder, S.E. & Emr, S.D. A multispecificity syntaxin homologue, Vam3p, essential for autophagic and biosynthetic protein transport to the vacuole. J Cell Biol 138, 517-529 (1997). 103. Fischer von Mollard, G. & Stevens, T.H. The Saccharomyces cerevisiae v-SNARE Vti1p is required for multiple membrane transport pathways to the vacuole. Mol Biol Cell 10, 1719-1732 (1999). 104. Harding, T.M., Morano, K.A., Scott, S.V. & Klionsky, D.J. Isolation and characterization of yeast mutants in the cytoplasm to vacuole protein targeting pathway. J Cell Biol 131, 591-602 (1995). 105. Sato, T.K., Darsow, T. & Emr, S.D. Vam7p, a SNAP-25-like molecule, and Vam3p, a syntaxin homolog, function together in yeast vacuolar protein trafficking. Mol Cell Biol 18, 5308-5319 (1998). 106. Wang, C.W., Stromhaug, P.E., Kauffman, E.J., Weisman, L.S. & Klionsky, D.J. Yeast homotypic vacuole fusion requires the Ccz1-Mon1 complex during the tethering/docking stage. J Cell Biol 163, 973-985 (2003). 107. Wang, C.W., Stromhaug, P.E., Shima, J. & Klionsky, D.J. The Ccz1-Mon1 protein complex is required for the late step of multiple vacuole delivery pathways. J Biol Chem 277, 47917-47927 (2002). 108. Jager, S. et al. Role for Rab7 in maturation of late autophagic vacuoles. J Cell Sci 117, 4837-4848 (2004). 109. Sanchez-Wandelmer, J. & Reggiori, F. Amphisomes: out of the autophagosome shadow? EMBO J 32, 3116- 3118 (2013). 110. Yang, Z., Huang, J., Geng, J., Nair, U. & Klionsky, D.J. Atg22 recycles amino acids to link the degradative and recycling functions of autophagy. Mol Biol Cell 17, 5094-5104 (2006). 111. Lynch-Day, M.A. & Klionsky, D.J. The Cvt pathway as a model for selective autophagy. FEBS Lett 584, 1359-1366 (2010). 112. Leber, R., Silles, E., Sandoval, I.V. & Mazon, M.J. Yol082p, a novel CVT protein involved in the selective targeting of aminopeptidase I to the yeast vacuole. J Biol Chem 276, 29210-29217 (2001). 113. Scott, S.V., Guan, J., Hutchins, M.U., Kim, J. & Klionsky, D.J. Cvt19 is a receptor for the cytoplasm-to- vacuole targeting pathway. Mol Cell 7, 1131-1141 (2001). 114. Shintani, T., Huang, W.P., Stromhaug, P.E. & Klionsky, D.J. Mechanism of cargo selection in the cytoplasm to vacuole targeting pathway. Dev Cell 3, 825-837 (2002). 115. Yorimitsu, T. & Klionsky, D.J. Atg11 links cargo to the vesicle-forming machinery in the cytoplasm to vacuole targeting pathway. Mol Biol Cell 16, 1593-1605 (2005). 116. Pfaffenwimmer, T. et al. Hrr25 kinase promotes selective autophagy by phosphorylating the cargo receptor Atg19. EMBO Rep 15, 862-870 (2014). 117. Xie, Z., Nair, U. & Klionsky, D.J. Atg8 controls phagophore expansion during autophagosome formation. Mol Biol Cell 19, 3290-3298 (2008). 118. Suzuki, K., Kondo, C., Morimoto, M. & Ohsumi, Y. Selective transport of alpha-mannosidase by autophagic pathways: identification of a novel receptor, Atg34p. J Biol Chem 285, 30019-30025 (2010). 119. Kanki, T., Wang, K., Cao, Y., Baba, M. & Klionsky, D.J. Atg32 is a mitochondrial protein that confers selectivity during mitophagy. Dev Cell 17, 98-109 (2009). 120. Farre, J.C., Manjithaya, R., Mathewson, R.D. & Subramani, S. PpAtg30 tags peroxisomes for turnover by selective autophagy. Dev Cell 14, 365-376 (2008). 121. Motley, A.M., Nuttall, J.M. & Hettema, E.H. Pex3-anchored Atg36 tags peroxisomes for degradation in Saccharomyces cerevisiae. EMBO J 31, 2852-2868 (2012). 122. Noda, N.N. et al. Structural basis of target recognition by Atg8/LC3 during selective autophagy. Genes Cells 13, 1211-1218 (2008). 123. Pankiv, S. et al. p62/SQSTM1 binds directly to Atg8/LC3 to facilitate degradation of ubiquitinated protein aggregates by autophagy. J Biol Chem 282, 24131-24145 (2007). 124. Klionsky, D.J. & Schulman, B.A. Dynamic regulation of macroautophagy by distinctive ubiquitin-like proteins. Nat Struct Mol Biol 21, 336-345 (2014). 125. Pickford, F. et al. The autophagy-related protein beclin 1 shows reduced expression in early Alzheimer disease and regulates amyloid beta accumulation in mice. J Clin Invest 118, 2190-2199 (2008). 126. Ravikumar, B. et al. Inhibition of mTOR induces autophagy and reduces toxicity of polyglutamine expansions in fly and mouse models of Huntington disease. Nat Genet 36, 585-595 (2004). 127. Winslow, A.R. et al. alpha-Synuclein impairs macroautophagy: implications for Parkinson's disease. J Cell Biol 190, 1023-1037 (2010). 128. Lu, K., Psakhye, I. & Jentsch, S. Autophagic clearance of polyQ proteins mediated by ubiquitin-Atg8 adaptors of the conserved CUET protein family. Cell 158, 549-563 (2014). 129. Lu, K., Psakhye, I. & Jentsch, S. A new class of ubiquitin-Atg8 receptors involved in selective autophagy and polyQ protein clearance. Autophagy 10, 2381-2382 (2014).

139 130. Kim, P.K., Hailey, D.W., Mullen, R.T. & Lippincott-Schwartz, J. Ubiquitin signals autophagic degradation of cytosolic proteins and peroxisomes. Proc Natl Acad Sci U S A 105, 20567-20574 (2008). 131. Kirkin, V. et al. A role for NBR1 in autophagosomal degradation of ubiquitinated substrates. Mol Cell 33, 505-516 (2009). 132. Shen, Z., Li, Y., Gasparski, A.N., Abeliovich, H. & Greenberg, M.L. Cardiolipin Regulates Mitophagy through the Protein Kinase C Pathway. J Biol Chem 292, 2916-2923 (2017). 133. Lamark, T., Kirkin, V., Dikic, I. & Johansen, T. NBR1 and p62 as cargo receptors for selective autophagy of ubiquitinated targets. Cell Cycle 8, 1986-1990 (2009). 134. Rogov, V.V. et al. Phosphorylation of the mitochondrial autophagy receptor Nix enhances its interaction with LC3 proteins. Sci Rep 7, 1131 (2017). 135. Filimonenko, M. et al. The selective macroautophagic degradation of aggregated proteins requires the PI3P- binding protein Alfy. Mol Cell 38, 265-279 (2010). 136. Clausen, T.H. et al. p62/SQSTM1 and ALFY interact to facilitate the formation of p62 bodies/ALIS and their degradation by autophagy. Autophagy 6, 330-344 (2010). 137. Lystad, A.H. et al. Structural determinants in GABARAP required for the selective binding and recruitment of ALFY to LC3B-positive structures. EMBO Rep 15, 557-565 (2014). 138. Simonsen, A. et al. Alfy, a novel FYVE-domain-containing protein associated with protein granules and autophagic membranes. J Cell Sci 117, 4239-4251 (2004). 139. Korolchuk, V.I., Menzies, F.M. & Rubinsztein, D.C. Mechanisms of cross-talk between the ubiquitin- proteasome and autophagy-lysosome systems. FEBS Lett 584, 1393-1398 (2010). 140. Verhoef, L.G., Lindsten, K., Masucci, M.G. & Dantuma, N.P. Aggregate formation inhibits proteasomal degradation of polyglutamine proteins. Hum Mol Genet 11, 2689-2700 (2002). 141. Lu, K., den Brave, F. & Jentsch, S. Receptor oligomerization guides pathway choice between proteasomal and autophagic degradation. Nat Cell Biol 19, 732-739 (2017). 142. Wurzer, B. et al. Oligomerization of p62 allows for selection of ubiquitinated cargo and isolation membrane during selective autophagy. Elife 4, e08941 (2015). 143. Klionsky, D.J. & Ohsumi, Y. Vacuolar import of proteins and organelles from the cytoplasm. Annu Rev Cell Dev Biol 15, 1-32 (1999). 144. Nazarko, T.Y., Farre, J.C. & Subramani, S. Peroxisome size provides insights into the function of autophagy- related proteins. Mol Biol Cell 20, 3828-3839 (2009). 145. Farre, J.C., Burkenroad, A., Burnett, S.F. & Subramani, S. Phosphorylation of mitophagy and pexophagy receptors coordinates their interaction with Atg8 and Atg11. EMBO Rep 14, 441-449 (2013). 146. Kim, J. et al. Cvt9/Gsa9 functions in sequestering selective cytosolic cargo destined for the vacuole. J Cell Biol 153, 381-396 (2001). 147. Nazarko, T.Y. et al. Peroxisomal Atg37 binds Atg30 or palmitoyl-CoA to regulate phagophore formation during pexophagy. J Cell Biol 204, 541-557 (2014). 148. Deosaran, E. et al. NBR1 acts as an autophagy receptor for peroxisomes. J Cell Sci 126, 939-952 (2013). 149. Yamashita, S., Abe, K., Tatemichi, Y. & Fujiki, Y. The membrane peroxin PEX3 induces peroxisome- ubiquitination-linked pexophagy. Autophagy 10, 1549-1564 (2014). 150. Nordgren, M. et al. Export-deficient monoubiquitinated PEX5 triggers peroxisome removal in SV40 large T antigen-transformed mouse embryonic fibroblasts. Autophagy 11, 1326-1340 (2015). 151. Zhang, J. et al. ATM functions at the peroxisome to induce pexophagy in response to ROS. Nat Cell Biol 17, 1259-1269 (2015). 152. Grou, C.P. et al. The peroxisomal protein import machinery--a case report of transient ubiquitination with a new flavor. Cell Mol Life Sci 66, 254-262 (2009). 153. Sargent, G. et al. PEX2 is the E3 ubiquitin ligase required for pexophagy during starvation. J Cell Biol 214, 677-690 (2016). 154. Hara-Kuge, S. & Fujiki, Y. The peroxin Pex14p is involved in LC3-dependent degradation of mammalian peroxisomes. Exp Cell Res 314, 3531-3541 (2008). 155. Liu, L., Sakakibara, K., Chen, Q. & Okamoto, K. Receptor-mediated mitophagy in yeast and mammalian systems. Cell Res 24, 787-795 (2014). 156. Wang, K., Jin, M., Liu, X. & Klionsky, D.J. Proteolytic processing of Atg32 by the mitochondrial i-AAA protease Yme1 regulates mitophagy. Autophagy 9, 1828-1836 (2013). 157. Redmann, M., Dodson, M., Boyer-Guittaut, M., Darley-Usmar, V. & Zhang, J. Mitophagy mechanisms and role in human diseases. Int J Biochem Cell Biol 53, 127-133 (2014).

140 158. Kurihara, Y. et al. Mitophagy plays an essential role in reducing mitochondrial production of reactive oxygen species and mutation of mitochondrial DNA by maintaining mitochondrial quantity and quality in yeast. J Biol Chem 287, 3265-3272 (2012). 159. Stotland, A. & Gottlieb, R.A. Mitochondrial quality control: Easy come, easy go. Biochim Biophys Acta 1853, 2802-2811 (2015). 160. Von Stockum, S., Nardin, A., Schrepfer, E. & Ziviani, E. Mitochondrial dynamics and mitophagy in Parkinson's disease: A fly point of view. Neurobiol Dis 90, 58-67 (2016). 161. Aihara, M. et al. Tor and the Sin3-Rpd3 complex regulate expression of the mitophagy receptor protein Atg32 in yeast. J Cell Sci 127, 3184-3196 (2014). 162. Kanki, T., Furukawa, K. & Yamashita, S. Mitophagy in yeast: Molecular mechanisms and physiological role. Biochim Biophys Acta 1853, 2756-2765 (2015). 163. Kanki, T. & Klionsky, D.J. Mitophagy in yeast occurs through a selective mechanism. J Biol Chem 283, 32386-32393 (2008). 164. Okamoto, K., Kondo-Okamoto, N. & Ohsumi, Y. Mitochondria-anchored receptor Atg32 mediates degradation of mitochondria via selective autophagy. Dev Cell 17, 87-97 (2009). 165. Mao, K., Wang, K., Liu, X. & Klionsky, D.J. The scaffold protein Atg11 recruits fission machinery to drive selective mitochondria degradation by autophagy. Dev Cell 26, 9-18 (2013). 166. Kondo-Okamoto, N. et al. Autophagy-related protein 32 acts as autophagic degron and directly initiates mitophagy. J Biol Chem 287, 10631-10638 (2012). 167. Aoki, Y. et al. Phosphorylation of Serine 114 on Atg32 mediates mitophagy. Mol Biol Cell 22, 3206-3217 (2011). 168. Kanki, T. et al. Casein kinase 2 is essential for mitophagy. EMBO Rep 14, 788-794 (2013). 169. Mao, K., Wang, K., Zhao, M., Xu, T. & Klionsky, D.J. Two MAPK-signaling pathways are required for mitophagy in Saccharomyces cerevisiae. J Cell Biol 193, 755-767 (2011). 170. Abeliovich, H., Zarei, M., Rigbolt, K.T., Youle, R.J. & Dengjel, J. Involvement of mitochondrial dynamics in the segregation of mitochondrial matrix proteins during stationary phase mitophagy. Nat Commun 4, 2789 (2013). 171. Georgakopoulos, N.D., Wells, G. & Campanella, M. The pharmacological regulation of cellular mitophagy. Nat Chem Biol 13, 136-146 (2017). 172. Narendra, D.P. et al. PINK1 is selectively stabilized on impaired mitochondria to activate Parkin. PLoS Biol 8, e1000298 (2010). 173. Sandoval, H. et al. Essential role for Nix in autophagic maturation of erythroid cells. Nature 454, 232-235 (2008). 174. Wei, Y., Chiang, W.C., Sumpter, R., Jr., Mishra, P. & Levine, B. Prohibitin 2 Is an Inner Mitochondrial Membrane Mitophagy Receptor. Cell 168, 224-238 e210 (2017). 175. Durcan, T.M. & Fon, E.A. The three 'P's of mitophagy: PARKIN, PINK1, and post-translational modifications. Genes Dev 29, 989-999 (2015). 176. Kondapalli, C. et al. PINK1 is activated by mitochondrial membrane potential depolarization and stimulates Parkin E3 ligase activity by phosphorylating Serine 65. Open Biol 2, 120080 (2012). 177. Trempe, J.F. et al. Structure of parkin reveals mechanisms for ubiquitin ligase activation. Science 340, 1451- 1455 (2013). 178. Yamano, K. & Youle, R.J. PINK1 is degraded through the N-end rule pathway. Autophagy 9, 1758-1769 (2013). 179. Aerts, L., Craessaerts, K., De Strooper, B. & Morais, V.A. PINK1 kinase catalytic activity is regulated by phosphorylation on serines 228 and 402. J Biol Chem 290, 2798-2811 (2015). 180. Gegg, M.E. et al. Mitofusin 1 and mitofusin 2 are ubiquitinated in a PINK1/parkin-dependent manner upon induction of mitophagy. Hum Mol Genet 19, 4861-4870 (2010). 181. Kane, L.A. et al. PINK1 phosphorylates ubiquitin to activate Parkin E3 ubiquitin ligase activity. J Cell Biol 205, 143-153 (2014). 182. Koyano, F. et al. Ubiquitin is phosphorylated by PINK1 to activate parkin. Nature 510, 162-166 (2014). 183. Chen, Y. & Dorn, G.W., 2nd PINK1-phosphorylated mitofusin 2 is a Parkin receptor for culling damaged mitochondria. Science 340, 471-475 (2013). 184. Pickrell, A.M. & Youle, R.J. The roles of PINK1, parkin, and mitochondrial fidelity in Parkinson's disease. Neuron 85, 257-273 (2015). 185. Geisler, S. et al. PINK1/Parkin-mediated mitophagy is dependent on VDAC1 and p62/SQSTM1. Nat Cell Biol 12, 119-131 (2010).

141 186. Lazarou, M. et al. The ubiquitin kinase PINK1 recruits autophagy receptors to induce mitophagy. Nature 524, 309-314 (2015). 187. Bingol, B. et al. The mitochondrial deubiquitinase USP30 opposes parkin-mediated mitophagy. Nature 510, 370-375 (2014). 188. Fang, L. et al. Mitochondrial function in neuronal cells depends on p97/VCP/Cdc48-mediated quality control. Front Cell Neurosci 9, 16 (2015). 189. McLelland, G.L., Lee, S.A., McBride, H.M. & Fon, E.A. Syntaxin-17 delivers PINK1/parkin-dependent mitochondrial vesicles to the endolysosomal system. J Cell Biol 214, 275-291 (2016). 190. Sugiura, A., McLelland, G.L., Fon, E.A. & McBride, H.M. A new pathway for mitochondrial quality control: mitochondrial-derived vesicles. EMBO J 33, 2142-2156 (2014). 191. Matsushima, M. et al. Isolation, mapping, and functional analysis of a novel human cDNA (BNIP3L) encoding a protein homologous to human NIP3. Genes Chromosomes Cancer 21, 230-235 (1998). 192. Wang, X. et al. PINK1 and Parkin target Miro for phosphorylation and degradation to arrest mitochondrial motility. Cell 147, 893-906 (2011). 193. Sheng, Z.H. & Cai, Q. Mitochondrial transport in neurons: impact on synaptic homeostasis and neurodegeneration. Nat Rev Neurosci 13, 77-93 (2012). 194. Novak, I. et al. Nix is a selective autophagy receptor for mitochondrial clearance. EMBO Rep 11, 45-51 (2010). 195. Hsieh, C.H. et al. Functional Impairment in Miro Degradation and Mitophagy Is a Shared Feature in Familial and Sporadic Parkinson's Disease. Cell Stem Cell 19, 709-724 (2016). 196. Novak, I. & Dikic, I. Autophagy receptors in developmental clearance of mitochondria. Autophagy 7, 301- 303 (2011). 197. Zhang, J. et al. A short linear motif in BNIP3L (NIX) mediates mitochondrial clearance in reticulocytes. Autophagy 8, 1325-1332 (2012). 198. Ni, H.M., Williams, J.A. & Ding, W.X. Mitochondrial dynamics and mitochondrial quality control. Redox Biol 4, 6-13 (2015). 199. Chourasia, A.H., Boland, M.L. & Macleod, K.F. Mitophagy and cancer. Cancer Metab 3, 4 (2015). 200. Liu, L. et al. Mitochondrial outer-membrane protein FUNDC1 mediates hypoxia-induced mitophagy in mammalian cells. Nat Cell Biol 14, 177-185 (2012). 201. Bordi, M., Nazio, F. & Campello, S. The Close Interconnection between Mitochondrial Dynamics and Mitophagy in Cancer. Front Oncol 7, 81 (2017). 202. Chen, G. et al. A regulatory signaling loop comprising the PGAM5 phosphatase and CK2 controls receptor- mediated mitophagy. Mol Cell 54, 362-377 (2014). 203. Palikaras, K., Lionaki, E. & Tavernarakis, N. Mitophagy: In sickness and in health. Mol Cell Oncol 3, e1056332 (2016). 204. Zhang, H. et al. Mitochondrial autophagy is an HIF-1-dependent adaptive metabolic response to hypoxia. J Biol Chem 283, 10892-10903 (2008). 205. Shirihai, O.S., Song, M. & Dorn, G.W., 2nd How mitochondrial dynamism orchestrates mitophagy. Circ Res 116, 1835-1849 (2015). 206. Chen, M. et al. Mitophagy receptor FUNDC1 regulates mitochondrial dynamics and mitophagy. Autophagy 12, 689-702 (2016). 207. Wu, W. et al. FUNDC1 regulates mitochondrial dynamics at the ER-mitochondrial contact site under hypoxic conditions. EMBO J 35, 1368-1384 (2016). 208. Chu, C.T. et al. Cardiolipin externalization to the outer mitochondrial membrane acts as an elimination signal for mitophagy in neuronal cells. Nat Cell Biol 15, 1197-1205 (2013). 209. Sentelle, R.D. et al. Ceramide targets autophagosomes to mitochondria and induces lethal mitophagy. Nat Chem Biol 8, 831-838 (2012). 210. Bravo, R. et al. Endoplasmic reticulum and the unfolded protein response: dynamics and metabolic integration. Int Rev Cell Mol Biol 301, 215-290 (2013). 211. Yorimitsu, T., Nair, U., Yang, Z. & Klionsky, D.J. Endoplasmic reticulum stress triggers autophagy. J Biol Chem 281, 30299-30304 (2006). 212. Bernales, S., McDonald, K.L. & Walter, P. Autophagy counterbalances endoplasmic reticulum expansion during the unfolded protein response. PLoS Biol 4, e423 (2006). 213. Schuck, S., Gallagher, C.M. & Walter, P. ER-phagy mediates selective degradation of endoplasmic reticulum independently of the core autophagy machinery. J Cell Sci 127, 4078-4088 (2014).

142 214. Khaminets, A. et al. Regulation of endoplasmic reticulum turnover by selective autophagy. Nature 522, 354- 358 (2015). 215. Mochida, K. et al. Receptor-mediated selective autophagy degrades the endoplasmic reticulum and the nucleus. Nature 522, 359-362 (2015). 216. Changou, C.A. et al. Arginine starvation-associated atypical cellular death involves mitochondrial dysfunction, nuclear DNA leakage, and chromatin autophagy. Proc Natl Acad Sci U S A 111, 14147-14152 (2014). 217. Dou, Z. et al. Autophagy mediates degradation of nuclear lamina. Nature 527, 105-109 (2015). 218. Mijaljica, D., Prescott, M. & Devenish, R.J. A late form of nucleophagy in Saccharomyces cerevisiae. PLoS One 7, e40013 (2012). 219. Kvam, E. & Goldfarb, D.S. Structure and function of nucleus-vacuole junctions: outer-nuclear-membrane targeting of Nvj1p and a role in tryptophan uptake. J Cell Sci 119, 3622-3633 (2006). 220. Kvam, E. & Goldfarb, D.S. Nucleus-vacuole junctions and piecemeal microautophagy of the nucleus in S. cerevisiae. Autophagy 3, 85-92 (2007). 221. Maejima, I. et al. Autophagy sequesters damaged lysosomes to control lysosomal biogenesis and kidney injury. EMBO J 32, 2336-2347 (2013). 222. Colombo, M.I., Gutierrez, M.G. & Romano, P.S. The two faces of autophagy: Coxiella and Mycobacterium. Autophagy 2, 162-164 (2006). 223. Gomes, L.C. & Dikic, I. Autophagy in antimicrobial immunity. Mol Cell 54, 224-233 (2014). 224. Zheng, Y.T. et al. The adaptor protein p62/SQSTM1 targets invading bacteria to the autophagy pathway. J Immunol 183, 5909-5916 (2009). 225. Thurston, T.L., Ryzhakov, G., Bloor, S., von Muhlinen, N. & Randow, F. The TBK1 adaptor and autophagy receptor NDP52 restricts the proliferation of ubiquitin-coated bacteria. Nat Immunol 10, 1215-1221 (2009). 226. Thurston, T.L., Wandel, M.P., von Muhlinen, N., Foeglein, A. & Randow, F. Galectin 8 targets damaged vesicles for autophagy to defend cells against bacterial invasion. Nature 482, 414-418 (2012). 227. Wild, P. et al. Phosphorylation of the autophagy receptor optineurin restricts Salmonella growth. Science 333, 228-233 (2011). 228. Singh, R. et al. Autophagy regulates lipid metabolism. Nature 458, 1131-1135 (2009). 229. van Zutphen, T. et al. Lipid droplet autophagy in the yeast Saccharomyces cerevisiae. Mol Biol Cell 25, 290- 301 (2014). 230. Kaur, J. & Debnath, J. Autophagy at the crossroads of catabolism and anabolism. Nat Rev Mol Cell Biol 16, 461-472 (2015). 231. Pantopoulos, K., Porwal, S.K., Tartakoff, A. & Devireddy, L. Mechanisms of mammalian iron homeostasis. Biochemistry 51, 5705-5724 (2012). 232. Asano, T. et al. Distinct mechanisms of ferritin delivery to lysosomes in iron-depleted and iron-replete cells. Mol Cell Biol 31, 2040-2052 (2011). 233. Mancias, J.D. et al. Ferritinophagy via NCOA4 is required for erythropoiesis and is regulated by iron dependent HERC2-mediated proteolysis. Elife 4 (2015). 234. Mancias, J.D., Wang, X., Gygi, S.P., Harper, J.W. & Kimmelman, A.C. Quantitative proteomics identifies NCOA4 as the cargo receptor mediating ferritinophagy. Nature 509, 105-109 (2014). 235. Delbridge, L.M., Mellor, K.M., Taylor, D.J. & Gottlieb, R.A. Myocardial autophagic energy stress responses- -macroautophagy, mitophagy, and glycophagy. Am J Physiol Heart Circ Physiol 308, H1194-1204 (2015). 236. Ueno, T. & Komatsu, M. Autophagy in the liver: functions in health and disease. Nat Rev Gastroenterol Hepatol 14, 170-184 (2017). 237. Zhu, Y., Zhang, M., Kelly, A.R. & Cheng, A. The carbohydrate-binding domain of overexpressed STBD1 is important for its stability and protein-protein interactions. Biosci Rep 34 (2014). 238. Hara, T. et al. Suppression of basal autophagy in neural cells causes neurodegenerative disease in mice. Nature 441, 885-889 (2006). 239. Feng, Y. et al. Phosphorylation of Atg9 regulates movement to the phagophore assembly site and the rate of autophagosome formation. Autophagy 12, 648-658 (2016). 240. Feng, Y., Yao, Z. & Klionsky, D.J. How to control self-digestion: transcriptional, post-transcriptional, and post-translational regulation of autophagy. Trends Cell Biol 25, 354-363 (2015). 241. Bernard, A. et al. Rph1/KDM4 mediates nutrient-limitation signaling that leads to the transcriptional induction of autophagy. Curr Biol 25, 546-555 (2015). 242. Bernard, A., Jin, M., Xu, Z. & Klionsky, D.J. A large-scale analysis of autophagy-related gene expression identifies new regulators of autophagy. Autophagy 11, 2114-2122 (2015).

143 243. Hu, G. et al. A conserved mechanism of TOR-dependent RCK-mediated mRNA degradation regulates autophagy. Nat Cell Biol 17, 930-942 (2015). 244. Garneau, N.L., Wilusz, J. & Wilusz, C.J. The highways and byways of mRNA decay. Nat Rev Mol Cell Biol 8, 113-126 (2007). 245. Parker, R. RNA degradation in Saccharomyces cerevisae. Genetics 191, 671-702 (2012). 246. Tharun, S. Lsm1-7-Pat1 complex: a link between 3' and 5'-ends in mRNA decay? RNA Biol 6, 228-232 (2009). 247. Bouveret, E., Rigaut, G., Shevchenko, A., Wilm, M. & Seraphin, B. A Sm-like protein complex that participates in mRNA degradation. EMBO J 19, 1661-1671 (2000). 248. Chowdhury, A., Kalurupalle, S. & Tharun, S. Pat1 contributes to the RNA binding activity of the Lsm1-7- Pat1 complex. RNA 20, 1465-1475 (2014). 249. Chowdhury, A., Mukhopadhyay, J. & Tharun, S. The decapping activator Lsm1p-7p-Pat1p complex has the intrinsic ability to distinguish between oligoadenylated and polyadenylated RNAs. RNA 13, 998-1016 (2007). 250. Chowdhury, A. & Tharun, S. Activation of decapping involves binding of the mRNA and facilitation of the post-binding steps by the Lsm1-7-Pat1 complex. RNA 15, 1837-1848 (2009). 251. He, W. & Parker, R. The yeast cytoplasmic LsmI/Pat1p complex protects mRNA 3' termini from partial degradation. Genetics 158, 1445-1455 (2001). 252. Tharun, S., Muhlrad, D., Chowdhury, A. & Parker, R. Mutations in the Saccharomyces cerevisiae LSM1 gene that affect mRNA decapping and 3' end protection. Genetics 170, 33-46 (2005). 253. Noda, T. & Klionsky, D.J. The quantitative Pho8Delta60 assay of nonspecific autophagy. Methods Enzymol 451, 33-42 (2008). 254. Shintani, T. & Klionsky, D.J. Cargo proteins facilitate the formation of transport vesicles in the cytoplasm to vacuole targeting pathway. J Biol Chem 279, 29889-29894 (2004). 255. Morawska, M. & Ulrich, H.D. An expanded tool kit for the auxin-inducible degron system in budding yeast. Yeast 30, 341-351 (2013). 256. Anderson, J.S. & Parker, R.P. The 3' to 5' degradation of yeast mRNAs is a general mechanism for mRNA turnover that requires the SKI2 DEVH box protein and 3' to 5' exonucleases of the exosome complex. EMBO J 17, 1497-1506 (1998). 257. van Hoof, A., Staples, R.R., Baker, R.E. & Parker, R. Function of the ski4p (Csl4p) and Ski7p proteins in 3'- to-5' degradation of mRNA. Mol Cell Biol 20, 8230-8243 (2000). 258. Brown, J.T., Bai, X. & Johnson, A.W. The yeast antiviral proteins Ski2p, Ski3p, and Ski8p exist as a complex in vivo. RNA 6, 449-457 (2000). 259. Araki, Y. et al. Ski7p G protein interacts with the exosome and the Ski complex for 3'-to-5' mRNA decay in yeast. EMBO J 20, 4684-4693 (2001). 260. Ramachandran, V., Shah, K.H. & Herman, P.K. The cAMP-dependent protein kinase signaling pathway is a key regulator of P body foci formation. Mol Cell 43, 973-981 (2011). 261. Shah, K.H., Zhang, B., Ramachandran, V. & Herman, P.K. Processing body and stress granule assembly occur by independent and differentially regulated pathways in Saccharomyces cerevisiae. Genetics 193, 109- 123 (2013). 262. Chowdhury, A. & Tharun, S. lsm1 mutations impairing the ability of the Lsm1p-7p-Pat1p complex to preferentially bind to oligoadenylated RNA affect mRNA decay in vivo. RNA 14, 2149-2158 (2008). 263. Selth, L.A., Gilbert, C. & Svejstrup, J.Q. RNA immunoprecipitation to determine RNA-protein associations in vivo. Cold Spring Harb Protoc 2009, pdb prot5234 (2009). 264. Hocine, S., Raymond, P., Zenklusen, D., Chao, J.A. & Singer, R.H. Single-molecule analysis of gene expression using two-color RNA labeling in live yeast. Nat Methods 10, 119-121 (2013). 265. Scheller, N. et al. Identification of PatL1, a human homolog to yeast P body component Pat1. Biochim Biophys Acta 1773, 1786-1792 (2007). 266. Ozgur, S., Chekulaeva, M. & Stoecklin, G. Human Pat1b connects deadenylation with mRNA decapping and controls the assembly of processing bodies. Mol Cell Biol 30, 4308-4323 (2010). 267. Delorme-Axford, E. et al. The exoribonuclease Xrn1 is a post-transcriptional regulator of autophagy. Autophagy in press (2018). 268. Song, M.G. & Kiledjian, M. 3' Terminal oligo U-tract-mediated stimulation of decapping. RNA 13, 2356- 2365 (2007). 269. Garre, E., Pelechano, V., Sanchez Del Pino, M., Alepuz, P. & Sunnerhagen, P. The Lsm1-7/Pat1 complex binds to stress-activated mRNAs and modulates the response to hyperosmotic shock. PLoS Genet 14, e1007563 (2018).

144 270. Nissan, T., Rajyaguru, P., She, M., Song, H. & Parker, R. Decapping activators in Saccharomyces cerevisiae act by multiple mechanisms. Mol Cell 39, 773-783 (2010). 271. Huber, A. et al. Characterization of the rapamycin-sensitive phosphoproteome reveals that Sch9 is a central coordinator of protein synthesis. Genes Dev 23, 1929-1943 (2009). 272. Stephan, J.S., Yeh, Y.Y., Ramachandran, V., Deminoff, S.J. & Herman, P.K. The Tor and cAMP-dependent protein kinase signaling pathways coordinately control autophagy in Saccharomyces cerevisiae. Autophagy 6, 294-295 (2010). 273. Schmelzle, T., Beck, T., Martin, D.E. & Hall, M.N. Activation of the RAS/cyclic AMP pathway suppresses a TOR deficiency in yeast. Mol Cell Biol 24, 338-351 (2004). 274. Soulard, A. et al. The rapamycin-sensitive phosphoproteome reveals that TOR controls protein kinase A toward some but not all substrates. Mol Biol Cell 21, 3475-3486 (2010). 275. Gueldener, U., Heinisch, J., Koehler, G.J., Voss, D. & Hegemann, J.H. A second set of loxP marker cassettes for Cre-mediated multiple gene knockouts in budding yeast. Nucleic Acids Res 30, e23 (2002). 276. Longtine, M.S. et al. Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14, 953-961 (1998). 277. Toulmay, A. & Schneiter, R. A two-step method for the introduction of single or multiple defined point mutations into the genome of Saccharomyces cerevisiae. Yeast 23, 825-831 (2006). 278. Li, C. et al. FastCloning: a highly simplified, purification-free, sequence- and ligation-independent PCR cloning method. BMC Biotechnol 11, 92 (2011). 279. Sung, M.K. & Huh, W.K. Bimolecular fluorescence complementation analysis system for in vivo detection of protein-protein interaction in Saccharomyces cerevisiae. Yeast 24, 767-775 (2007). 280. Swaney, D.L., McAlister, G.C. & Coon, J.J. Decision tree-driven tandem mass spectrometry for shotgun proteomics. Nat Methods 5, 959-964 (2008). 281. Delorme-Axford, E. et al. The exoribonuclease Xrn1 is a post-transcriptional negative regulator of autophagy. Autophagy 14, 898-912 (2018). 282. Wen, X. & Klionsky, D. An overview of macroautophagy in yeast. Journal of Molecular Biology 428, 1681- 1699 (2016). 283. Rao, Y., Perna, M.G., Hofmann, B., Beier, V. & Wollert, T. The Atg1–kinase complex tethers Atg9-vesicles to initiate autophagy. 284. Sanchez-Wandelmer, J. et al. Atg4 proteolytic activity can be inhibited by Atg1 phosphorylation. Nature Communications 8 (2017). 285. Ragusa, M., Stanley, R. & Hurley, J. Architecture of the Atg17 Complex as a Scaffold for Autophagosome Biogenesis. Cell 151, 1501-1512 (2012). 286. Chew, L. et al. Molecular interactions of the Saccharomyces cerevisiae Atg1 complex provide insights into assembly and regulatory mechanisms. Autophagy 11, 891-905 (2015). 287. Popelka, H. & Klionsky, D. The molecular mechanism of Atg13 function in autophagy induction: What is hidden behind the data? Autophagy 13, 449-451 (2017). 288. Yamamoto, H. et al. The Intrinsically Disordered Protein Atg13 Mediates Supramolecular Assembly of Autophagy Initiation Complexes. Developmental Cell 38, 86-99 (2016). 289. Stjepanovic, G. et al. Assembly and dynamics of the autophagy-initiating Atg1 complex. Proc Natl Acad Sci U S A 111, 12793-12798 (2014). 290. Kamada, Y. et al. Tor-mediated induction of autophagy via an Apg1 protein kinase complex. Journal of Cell Biology 150, 1507-1513 (2000). 291. Suzuki, S. et al. Atg13 HORMA domain recruits Atg9 vesicles during autophagosome formation. Proceedings of the National Academy of Sciences of the United States of America 112, 3350-3355 (2015). 292. Jao, C., Ragusa, M., Stanley, R. & Hurley, J. A HORMA domain in Atg13 mediates PI 3-kinase recruitment in autophagy. Proceedings of the National Academy of Sciences of the United States of America 110, 5486- 5491 (2013). 293. Fujioka, Y. et al. Structural basis of starvation-induced assembly of the autophagy initiation complex. Nat Struct Mol Biol 21, 513-521 (2014). 294. Scott, S. et al. Apg13p and Vac8p are part of a complex of phosphoproteins that are required for cytoplasm to vacuole targeting. Journal of Biological Chemistry 275, 25840-25849 (2000). 295. Yamamoto, H. et al. The Intrinsically Disordered Protein Atg13 Mediates Supramolecular Assembly of Autophagy Initiation Complexes. Dev Cell 38, 86-99 (2016). 296. Busch, D.J. et al. Intrinsically disordered proteins drive membrane curvature. Nat Commun 6, 7875 (2015).

145 297. Bartels, T. et al. The N-terminus of the intrinsically disordered protein alpha-synuclein triggers membrane binding and helix folding. Biophys J 99, 2116-2124 (2010). 298. Jeong, H. et al. Mechanistic insight into the nucleus-vacuole junction based on the Vac8p-Nvj1p crystal structure. Proc Natl Acad Sci U S A 114, E4539-E4548 (2017). 299. Roberts, P. et al. Piecemeal microautophagy of nucleus in Saccharomyces cerevisiae. Mol Biol Cell 14, 129- 141 (2003). 300. Cao, Y., Cheong, H., Song, H. & Klionsky, D.J. In vivo reconstitution of autophagy in Saccharomyces cerevisiae. J Cell Biol 182, 703-713 (2008). 301. Patil, A. & Nakamura, H. Disordered domains and high surface charge confer hubs with the ability to interact with multiple proteins in interaction networks. FEBS Lett 580, 2041-2045 (2006). 302. Habchi, J., Tompa, P., Longhi, S. & Uversky, V.N. Introducing protein intrinsic disorder. Chem Rev 114, 6561-6588 (2014). 303. Scott, S.V. et al. Apg13p and Vac8p are part of a complex of phosphoproteins that are required for cytoplasm to vacuole targeting. J Biol Chem 275, 25840-25849 (2000). 304. Gibson, T.J. Cell regulation: determined to signal discrete cooperation. Trends Biochem Sci 34, 471-482 (2009). 305. Bajaj, R., Bollen, M., Peti, W. & Page, R. KNL1 Binding to PP1 and Microtubules Is Mutually Exclusive. Structure 26, 1327-1336 e1324 (2018). 306. Noda, T. & Klionsky, D. THE QUANTITATIVE PHO8 Delta 60 ASSAY OF NONSPECIFIC AUTOPHAGY. Autophagy: Lower Eukaryotes and Non-Mammalian Systems, Pt a 451, 33-42 (2008). 307. Gatica, D. et al. The Pat1-Lsm Complex Stabilizes ATG mRNA during Nitrogen Starvation-Induced Autophagy. Mol Cell 73, 314-324 e314 (2019). 308. Hall, T.A. (Nucleic Acids Symposium, 1999).

146