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Record for a UF thesis. Title & abstract won't display until thesis is accessible after 2009-12-31.

Permanent Link: http://ufdc.ufl.edu/UFE0021725/00001

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Title: Record for a UF thesis. Title & abstract won't display until thesis is accessible after 2009-12-31.
Physical Description: Book
Language: english
Creator: Schroder, Laura Aaron
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2007

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Subjects / Keywords: Molecular Cell Biology (IDP) -- Dissertations, Academic -- UF
Genre: Medical Sciences thesis, Ph.D.
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theses   ( marcgt )
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Statement of Responsibility: by Laura Aaron Schroder.
Thesis: Thesis (Ph.D.)--University of Florida, 2007.
Local: Adviser: Dunn, William A.
Electronic Access: INACCESSIBLE UNTIL 2009-12-31

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Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2007
System ID: UFE0021725:00001

Permanent Link: http://ufdc.ufl.edu/UFE0021725/00001

Material Information

Title: Record for a UF thesis. Title & abstract won't display until thesis is accessible after 2009-12-31.
Physical Description: Book
Language: english
Creator: Schroder, Laura Aaron
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2007

Subjects

Subjects / Keywords: Molecular Cell Biology (IDP) -- Dissertations, Academic -- UF
Genre: Medical Sciences thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Statement of Responsibility: by Laura Aaron Schroder.
Thesis: Thesis (Ph.D.)--University of Florida, 2007.
Local: Adviser: Dunn, William A.
Electronic Access: INACCESSIBLE UNTIL 2009-12-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2007
System ID: UFE0021725:00001


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CHARACTERIZING THE ASSEMBLY OF THE MEMBRANES THAT SEQUESTER PEROXISOMES DURING AUTOPHAGY By LAURA A. SCHRODER A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2007 1

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2007 Laura A. Schroder 2

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This work was completed in remembrance of my dear brother, Glenn Eldon Miller, III, whom I love and miss dearly. You were here when I be gan this adventure; what would I give that you were here now! I sincerely hope that someday my work and the work of other scientists may curtail the suffering caused by metastic malignant melanoma and other cancers. This work is dedicated to my daughter, Mya Cata lina Schroder, who was born in the second year of the work that cumulated in my doctoral dissertation. Mya is th e light around which my universe gravitates: I cannot imagine my life without you. You endow those who are blessed with your presence immense joy. Recently you ha ve asked me what is meant by hope and peaceboth of which are embodied in the gift God gave us when he gave us you. This work could not have been accomplished without my long-suffering Mom, Lorraine Abate Miller. Your sheer determination to become a botanist inspired me in turn to become a molecular biologist, and your forbearance mode led my stubborn stick-to-it-ness. Thank you for instilling in me the importance of an educat ion. I truly appreciate all the sacrifices you have made and continue to make for your family. You have found solace in caring for Mya, and your hours are well spent caring for her during my ev enings and weekends in the laboratory. I truly value the support of Glenn Eldon Miller, Jr. and Linda Miller. You love me without limits, you cherish me for being me, and you do not judge. I am grateful to Jeremiah John Schroder for your loving support. Your patience and forbearance do not go unnoticed; I know you always try your be st. Your pride in this dissertation is appreciated. Our genetic experiment is more successful than I could have imagined. I will not forget the care and patience on the part of Ivonne Ulloa-Ron, Markus Patricio Tellkamp, and Daniela and Domenica Ulloa duri ng Myas time with you. Myas sweet, caring and loving personality and her amazing intelligence we re formed in her very tender first year and a half in part by all of you. The emotional and psychological support of Jolene Smith and Donna Tackett have kept me sane. That all friendships were so steadfast! You put things in perspective when times are hard (as they often are), and you always make me laugh. An extra special thanks to my professor W illiam A. Dunn, Jr., Ph.D. I cannot thank you enough for bestowing a quiet confidence in me with your initial instruction at the bench, your never failing to complement my early work, and your encouragement when science, by definition, is frustrating. You have challenged me to faithfully complete this task. I have learned to think and write scientifically because of you. I va lue your input, and your honesty astounds me. 3

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ACKNOWLEDGMENTS I am very grateful to my committee members John Aris, Ph.D., Brian Burke, Ph.D. and David Julian, Ph.D. for their helpful discussions, encour agement and help with editing. I especially thank William Dunn, Jr., Ph.D. for analyzing my wr iting with a fine-toothed comb. This work was supported by grants from the National Canc er Institute (NIH) CA95552 to W.A.Dunn, Jr., Ph.D. Specific Acknowledgments Chapter 2: This work was supported by NSF (MCB-9817002) and NIH (CA95552) grants to W.A. Dunn, Jr. and NIH (GM61156) grant to B. S. Glick. Chapter 3: This work was supported by grants from the National Sc ience Foundation Grant MCB-9817002 and the National Cancer Institute (NIH) CA95552 to W.A.Dunn, Jr. and a grant from The Norwegian Cancer Society to P. E. St rmhaug. We would like to thank Dr. B.S. Glick (University of Chicago) for ge nerously providing the pREMI-Z, pIB1, and pIB2 vectors. Finally, we would like to thank Mr. Todd Barnash for his help in assembling the figures, and Dr. Michelle Fry and Debbie Akin for helpful discussions and editing. Chapter 6: This work was supported by grants from the National Cancer Institute (NIH) CA95552 to W.A.Dunn, Jr. We woul d like to thank Dr. B. S. Glick (University of Chicago) for generously providing the pSar1T 34N, pSar1H79G, and pIB2 vectors and Dr. R.Y. Tsein for providing the mRFP gene. Finally, we would like to thank Mr. Todd Barnash for his help in assembling the figures, and Debbie Akin for helpful discussions and editing. I alone remain responsible for the content of the following, including any errors or omissions which may unwittingly remain. 4

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TABLE OF CONTENTS page ACKNOWLEDGMENTS...............................................................................................................4 LIST OF TABLES................................................................................................................. ..........9 LIST OF FIGURES.......................................................................................................................10 LIST OF TERMS...........................................................................................................................12 ABSTRACT...................................................................................................................................13 CHAPTER 1 INTRODUCTION..................................................................................................................15 The Role Of Atg9 In Glucose-Induced Mmicropexophagy...................................................18 Cellular Compartments Of Atg9.............................................................................................20 Transport Of Atg9 During Micrope xophagy Requires Vps15 And Atg11............................21 Vps15, A Component Of The Phosphatidyl Inositol-3 Kinase (Pi3k) Complex Thought To Signal The Start Of Autopha gy, Is Required For Atg9 Trafficking........21 Atg11 Is A Membrane-Associated Prot ein Required For Pexophagy But Not Autophagy....................................................................................................................25 Actin Is Required For Efficient Autophagy....................................................................28 The Atg1 Protein Kinase Is Re quired For Pexophagy And Autophagy..........................29 Atg9 Retrograde Trafficking Requires Atg23 And Atg27..............................................31 Components Of The Copii Secretory Pathway Are Required For Autophagy.......................32 Sar1 And Protein Transport.............................................................................................33 Sar1 And Lipid Transport................................................................................................35 Summation..............................................................................................................................35 2 IDENTIFICATION OF PEXOPHAGY GE NES BY RESTRICTION ENZYMEMEDIATED INTEGRATION (REMI)..................................................................................42 Introduction................................................................................................................... ..........42 Materials.................................................................................................................................43 Pichia Pastoris And E. Coli Strains.................................................................................43 Culture Media..................................................................................................................43 Vector..............................................................................................................................44 Transformation Of Pichia Pastoris .................................................................................44 Qualitative Assessment Of Alcohol Oxidase Degradation By Direct Colony Assay.....44 Quantitative Assessment Of AOX Degrad ation By Liquid Medium Assay...................44 Isolation Of Yeast Genomic DNA..................................................................................45 Methods..................................................................................................................................45 1.0. Vector Construction..................................................................................................46 2.0 Yeast Transformation................................................................................................46 5

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2.1 Electro-Competent Yeast...................................................................................46 2.2 Transformation Of Yeast By Electroporation....................................................47 2.3 Selection Of Transformants...............................................................................47 2.4 Verifying Vector Integration..............................................................................47 3.0. Identification And Isolation Of Glucose-Induced Pexophagy Mutants...................48 3.1 Direct Colony Assay..........................................................................................48 3.2 AOX Degradation By Liquid Medium Assay....................................................49 4.0 Identification Of The Disr upted Gene Caused By The Insertion Of Premi-Z..........50 4.1 Genomic DNA Isolation.....................................................................................50 4.2 Amplification Of Premi-Z Is olated From Pexophagy Mutants..........................51 4.3 Sequencing Flanking Genomic DNA.................................................................53 Notes.......................................................................................................................................53 3 PPATG9 TRAFFICKING DURING PEXOPHAGY............................................................61 Introduction................................................................................................................... ..........61 Experimental Procedures........................................................................................................ 64 Yeast Strains And Media.................................................................................................64 Yeast Transformation......................................................................................................64 Isolation Of GSA Mutants And Cloning Of GSA Genes By Restriction Enzyme Mediated Integration (REMI) Mutagenesis.................................................................65 Measurements Of Alcohol Oxidase (AOX) And Endogenous Protein Degradation......66 Western Blot Analysis.....................................................................................................67 Construction Of Ppatg Expression Vectors.....................................................................67 Fluorescence Microscopy And FM 4-64 Labeling..........................................................71 Electron Microscopy.......................................................................................................71 Results.....................................................................................................................................72 Ppatg9 Mutants Are Defective In Gl ucose-Induced Pexophagy And StarvationInduced Autophagy......................................................................................................72 Ppatg9 Is Essential For A Sequestration Event In Pexophagy........................................73 Cellular Localization Of Ppatg9 In Growing Cells.........................................................75 Cellular Trafficking Of Ppatg9 Du ring Glucose-Induced Pexophagy............................75 Cellular Trafficking Of Ppatg9 During Gl ucose-Induced Pexophagy Requires Other Ppatg Proteins..............................................................................................................77 Domains Of Ppatg9 Required Fo r Function And Trafficking..................................78 Perivacuolar Structures Contain Ppatg9 And Ppatg11, But Not Ppatg2.........................79 Ppatg9 Does Not Localize To MIPA Or The Pexophagosome.......................................80 Discussion...............................................................................................................................80 4 PPATG9 TRAFFICKING DURING MICROPEXOPHAGY IN PICHIA PASTORIS .......102 5 TARGETING MOTIFS OF ATG9......................................................................................107 Introduction................................................................................................................... ........107 Materials And Methods........................................................................................................10 9 Yeast Strains And Media...............................................................................................109 Yeast Transformation....................................................................................................110 6

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Isolation Of Gsa Mutants And Cloning Of GSA Genes By Restriction Enzyme Mediated Integration (REMI) Mutagenesis...............................................................110 Construction Of Ppatg9 Expression Vectors.................................................................111 Qualitative And Quantitative Assessment Of Alcohol Oxidase (AOX) Degradation..113 Western Blot Analysis...................................................................................................114 Measurements Of Protein Degradation.........................................................................115 Fluorescence Microscopy And FM 4-64 Labeling........................................................116 Results...................................................................................................................................116 Atg9 And Atg17 Occasionally Co-Localize..................................................................116 Ppatg9 Partial Deletion And Site-Specifi c Mutants Are Defective In GlucoseInduced Pexophagy And Starvation-Induced Autophagy..........................................116 Atg9 L3 And Atg9 579-668 Mutants Remain In The ER..........................................118 Atg9 QQHKA Remains At The PC9...........................................................................120 Atg9-W607A/Y611A Fails To Localized To The Sequestering Membranes (SM) On The Vacuole.........................................................................................................121 Atg9 N Localizes To The Vacuole But Is Not Functional By AOX Assay.................121 Sites Of Delay In Micropexophagy Caused By The Atg9 Mutants..............................122 Discussion.............................................................................................................................122 ER Exiting Motifs Of Atg9...........................................................................................123 PC9 Exiting Motif Of Atg9...........................................................................................124 PVS Exiting Motif Of Atg9...........................................................................................126 Functional Domain Of Atg9..........................................................................................127 Summary...............................................................................................................................128 6 SAR1P AND PEXOPHAGY..............................................................................................143 Introduction................................................................................................................... ........143 Materials And Methods........................................................................................................14 7 Yeast Strains And Media...............................................................................................147 Quantitative Assessment Of Alcohol Oxidase (AOX) Degradation.............................147 Measurements Of Protein Degradation.........................................................................148 Western Blot Analysis...................................................................................................148 Construction Of Conditional Expression Vectors.........................................................149 Yeast Transformation....................................................................................................150 Fluorescence Microscopy..............................................................................................150 Electron Microscopy.....................................................................................................150 Results...................................................................................................................................151 Effects Of Dominant-Neg ative Mutants Of Sar1p On Autophagy And Pexophagy.....151 Suppression Of Glucose-Induced Micr opexophagy By Dominant-Negative Mutants Of Sar1p Occurs At A Late Sequestration Event.......................................................153 Sar1p Is Essential For The Formation Of The MIPA....................................................154 The Lipidation Of Atg8p During Glucos e-Induced Micropex ophagy Is Altered Upon Expression Of Dominant -Negative Sar1p Mutants.........................................157 Sar1p Is Required For Early And Late Events Of Macropexophagy............................159 RFP-HDEL Localizes To Pexophagosomes When Sar1pH79G Is Expressed.............160 Discussion.............................................................................................................................160 Role Of Sar1p In Micropexophagy...............................................................................161 7

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8 Role Of Sar1p In Macropexophagy...............................................................................163 Summary...............................................................................................................................165 7 CONCLUSION.................................................................................................................. ...179 LIST OF REFERENCES.............................................................................................................181 BIOGRAPHICAL SKETCH.......................................................................................................190

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LIST OF TABLES Table page 1-1 Autophagy-related Genes.................................................................................................... ....37 1-2 Secretory-related Genes...........................................................................................................38 2-1 Pexophagy genes identify by REMI........................................................................................57 3-1 Pichia pastoris strains.............................................................................................................87 6-1 Pichia pastoris strains...........................................................................................................166 9

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LIST OF FIGURES Figure page 1-1 Peroxisome engulfment by glucose-induced micropexophagy...............................................39 1-2 Molecular events of the expansion of the sequestering membranes that engulf peroxisomes during micropexophagy................................................................................41 2-1 Insertional mutagenesis by restriction enzyme-mediated integration of linearized pREMI-Z............................................................................................................................58 2-2 Insertion of pREMI-Z into th e genomic DNA of REMI mutants...........................................59 2-3 The direct colony assay of AOX activity in REMI-mutated cells...........................................59 2-4 Glucose-induced degradation of AOX in REMI mutants.......................................................60 3-1 Pexophagy and autophagy are de fective in Ppatg9 mutants....................................................89 3-2 Pexophagy is blocked at an early sequestration event in cells lacking PpAtg9......................90 3-3 Cellular localiz ation of PpAtg9............................................................................................ ...91 3-4 PpAtg9 relocates from peripheral struct ures to the sequestering membranes during glucose-induced pexophagy...............................................................................................92 3-5 Trafficking of PpAtg9 in mutants def ective in early sequestration events..............................94 3-6 Trafficking of PpAtg9 in mutants defective in intermediate sequestration events..................95 3-7 Trafficking of PpAtg9 in mutants de fective in late sequestration events................................96 3-8 PpAtg9 domains required fo r function and trafficking...........................................................97 3-9 Perivacuolar structures contain PpAtg9 and PpAtg11............................................................98 3-10 PpAtg9 does not colocalize with PpAtg8 structures.............................................................99 3-11 Model of PpAtg9 tra fficking during pexophagy.................................................................101 4-1 Alignment of the putativ e ER exit motifs of Atg9................................................................105 4-2 Trafficking of PpAtg9 and other Atg proteins during micropexophagy...............................106 5-1 Localization of Atg9, Atg11 and Atg17 to the PVS.............................................................130 5-2 Linear structure of the PpAtg9 transmembr ane protein and deletions performed by sitedirected mutagenesis........................................................................................................132 10

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11 5-3 Atg9 mutants fail to suppor t glucose-induced micropexophagy...........................................133 5-4 Protein degradation of Atg9 muta nt cells during nitrogen starvation...................................134 5-5 GFP-atg9 L3 remains in the ER during glucose-induced micropexophagy........................135 5-6 GFP-atg9 489-502 and GFP-atg9 E491,4A remain in the ER during glucose-induced micropexophagy...............................................................................................................136 5-7 GFP-atg9 579-668 remains in the ER during glucose-induced micropexophagy................137 5-8 GFP-atg9 489-502, GFP-atg9 E491,4A, and GFP-atg9 579-668 localized to the nuclear membranes..........................................................................................................138 5-9 GFP-atg9 QQHKA remains at peripheral vesi cles during glucose-induced micropexophagy...............................................................................................................139 5-10 Distribution of Atg9 vesicl es on sucrose gradients.............................................................140 5-11 GFP-atg9(W607A/Y611A) fails to locali ze to the sequestering membranes (SM) during glucose-induced micropexophagy........................................................................141 5-12 GFP-Atg9, not GFP-atg9(W607A/Y611A), migrates as a doublet on Westerns................142 6-1 Sar1p is essential for autophagy and pexophagy...................................................................167 6-2 The expression of Sar1pT34N or Sar1pH79G suppresses a late sequestration event of micropexophagy...............................................................................................................168 6-3 The trafficking of Atg11p, Atg2p and Atg9p........................................................................169 6-4 Sar1p is essential for the formation of the MIPA..................................................................170 6-5 The effects of Sar1p on the lipidation of Atg8p....................................................................171 6-6 Effects of Sar1pT34N and Sar 1pH79G on ethanol-induced macropexophagy.....................172 6-7 Sar1pT34N suppresses formation of the pexophagosome ...................................................173 6-8 Components of the endoplasmic reticulum...........................................................................174 6-9 Model of the role of Sar1p in macropexophagy....................................................................175 6-10 Effects of Sar1pT34N and Sar1pH79G on cell growth.......................................................176 6-11 Stages of glucose-induced micropexophagy.......................................................................177 6-12 Cellular lo calization of Atg18p and Atg17p........................................................................178

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LIST OF TERMS Autophagy Derived from the Latin meaning self-eating. The process by which an organism sequesters and degrades superf luous organelles ut ilizing the lytic compartment MIPA Micropexophagic-specific membra ne apparatus which tethers the SM whereby homotypic fusion pr oceeds. The final expansion step cumulating in completion. Nucleation Expansion, completion se quential steps whereby micropexophagy proceeds phenotypically PAS Pre-autophagic structure adjacen t to the tips of the sequestering membranes PC9 Peripheral compartment in which Atg9p resides Pexophagy A subset of autophagy specific to methylotropic yeasts whereby the peroxisomes are selectively sequestered and degraded PVS Perivacuolar structure present at the base of the sequestering membranes SM Sequestering membranes; armlike extensions of the vacuole characterizing glucose-induced micropexophagy 12

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Abstract of Dissertation Pres ented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy CHARACTERIZING THE ASSEMBLY OF THE MEMBRANES THAT SEQUESTER PEROXISOMES DURING AUTOPHAGY By Laura A. Schroder December 2007 Chair: William A. Dunn, Jr. Major: Medical SciencesMoclecular Cell Biology The methylotropic yeast Pichia pastoris efficiently sequesters and degrades peroxisomes by micropexophagy when more easily degraded car bon sources become available. The process by which this degradation occurs during gluc ose-induced micropexophagy can be phenotypically characterized. Nucleation is the budding of the sequestering membranes (SM) from the vacuole. Expansion follows as the SM surround the peroxisomes. The micropexophagic-specific membrane apparatus (MIPA) bridges the opposing SM. Completion is the final enveloping of the peroxisomes for degrada tion within the vacuole. This study has characterized the molecular ev ents of SM expansion mediated by certain Atg proteins. PpAtg9p is an integral membrane pr otein essential for the ev ents of SM expansion. Upon synthesis, Atg9 transits from the endoplasmic reticulum (ER) to a peripheral Atg9 compartment (PC9). Furthermore, site-directed mutagenesis has identified two domains within Atg9 are essential for ER exit. Specifically, two critical amino acids have been identified that are essential for ER exit. The PC9 is unique exhibiting a density not shared by the ER, the mitochondria, and the vacuole. During gluc ose-induced micropexophagy, Atg9p transits through perivacuolar structures (PVS) at the base of the sequesteri ng membranes to the SM. The trafficking of Atg9 requires Atg11, Vps15, Atg2, and Atg7. The C-terminal QQHKA motif is 13

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14 required for exit of Atg9 from the PC9. Next, W607 and Y611 are essentia l for Atg9 transit to the SM. None of the Atg9 mutants is functional, including the N-terminal deletion that localizes properly to the SM. Thus, a number of amino acid motifs necessary for Atg9p trafficking have been identified and characterized. The role of Sar1p, a GTPase involved in COPII-mediated ER trafficking, in pexophagy was examined by expressing sar1(T34N) or sa r1(H79G) dominant negative mutants. By expressing these mutants, we have demonstrated that the lipidation of At g8 and the assembly of the micropexophagic-specific membrane apparatus (MIPA) require Sar1. Sar1 is also required for macropexopha gy. Pexophagosomes do not form in the sar1(T34N) mutant. Completion and/or fusi on of the pexophagosome with the vacuole is suppressed in the sar1(H79G) mutant Moreover, sar1(H79G) inhibits the retrograde transport of membranes containing HDEL, whic h supports the idea that autophagosome membranes originate in the endoplasmic reticulum. Our data demonstrate that the expansion of the SM during micropexophagy proceeds in two stages, an early Atg9-dependent that requires the PVS to expand the SM and a late Atg8pdependent that requires Sar1p and the PAS to assemble the MIPA.

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CHAPTER 1 INTRODUCTION Autophagy is the catabolic process by which the cytoplasm and the organelles therewith are sequestered into double-membrane organelles, transported to and degraded within the acidic lysosome or vacuole for possible recycling. Au tophagy may be non-selective or selective. An example of selective autophagy is mitophagy, th e major route by which the mitochondria are degraded (Abeliovich, 2007). Pexophagy is the se lective degradation of the peroxisomes by the vacuole in methylotropic yeasts such as Pichia pastoris (Dunn et al., 2005). Membranes of unknown origin condense around individual pe roxisomes forming double-membrane bound pexophagosomes which then fuse with the va cuole in the process of macropexophagy (Sakai et al. 2006). Clusters of peroxisomes are incorpor ated by sequestering membranes derived from the vacuole in the process of micropexophagy (Sakai et al. 2006). Micropexophagy proceeds by a specific set of phenotypic events characterized by nucleation, expansion and completion. Nucleati on, the initial budding of the vacuole to form structures reminiscent of the sequestering memb ranes (SM), is dependent on signal transduction events and does not require de novo protein synthesis or addi tional membrane sources. Expansion, the exaggeration of the SM to surr ound the peroxisomes, does require additional membranes and is thought to involve the form ation of large cisternal sheets by combining vesicles (Klionsky et al., 2007). Expansion can be divided into early expansion and late expansion in our experimental system. Membrane delivery during the late expansion event in micropexophagy is fulfilled by the micropexophag ic-specific membrane apparatus (MIPA), which conjoins the SM (Dunn et al., 2005). Completion occurs as th e SM fuse together and the peroxisomes are entirely surrounded by the vacuol e for degradation, duri ng which the double membranes are degraded for access to the peroxisomes inside. 15

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Glucose selective autophagy ( GSA) genes were originally identified based on the inability of gsa mutants to degrade the peroxisomes. A more straightforward shotgun approach as compared to a series of tedious yeast mating was desired, cumulating in Restriction EnzymeMediated Integration (REMI) whic h randomly integrates a vector containing the Zeocin cassette into the yeast genome (Schroder et al. 2007). Subsequent screening has enabled the identification of Gsa gene pr oducts. A unified yeast nomencl ature was developed as some proteins affect more than one autophagic pathway (Klionsky et al. 2003). Thus, the Apg, Aut, Cvt, Gsa, Paz and Pdd prefixes were dropped for the ATG abbreviation meaning a ut opha g yrelated genes (Table 1-1). Pexophagy is best studied in Pichia pastoris. P. pastoris is easily manipulated genetically (mutations are induced and proteins can be readily upregulated), molecularly (feedback from a dietetic switch results in the ra pid degradation of orga nelles) and biologically (the peroxisome cluster and the v acuole, which is larger than the nucleus, are visible on the light microscope). P. pastoris offers a unique perspective for an increasingly understood cellular phenomenonpexophagy. Micropexophagy is hereby di ssected into discrete genetic and morphologic steps. Pexophagy can be describe d genotypically by defining the ATG gene products required for signaling/recognition, expa nsion (nucleation, expansion and completion) and degradation (Figure 1-1). Signaling and recognition requ ire Pfk1, Vps15 and Atg11 (Yuan et al. 1997; Kim et al., 2001; Dunn et al., 2005). Nucleation requi res Atg1 (Abeliovich et al., 2000), Atg6 (Levine and Klionsky, 2004) Atg18, Vac8, and Vps15 (Stasyk et al., 1999; Guan et al. 2001; Fry et al., 2006). Expansion requires At g2, Atg7, Atg8, Atg9, and Atg11 (Chang et al. 2005). Completion requires Vac8 (Fry et al. 2006), Atg8, and Atg24 (Ano et al., 2005). Finally, degradation requires Pep4 and Prb1 (Tuttle and Dunn, 1995; Dunn et al., 2005). 16

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The expansion of the sequestering membranes (SM) is classified into early and late events. During the early events, the SM expands from perivacuolar struct ures (PVS). The late events are characterized by the assembly of the micropexophagy apparatus (MIPA) from the preautophagosome structure (PAS). Atg9 is a tran smembrane protein involved in both early and late expansion events. During micropexophagy, Atg9 traffics from a peripheral compartment to the PVS and the SM and to the PAS position at the tips of the SM. Atg11 is found at the PVS and SM, while Atg8 is at the PAS and MIPA (Figure 1-2). Membrane expansion during pexophagy is a rate-limiting process during pexophagy. Atg9 is the sole transmembrane protein shown to be required for the membrane trafficking events leading to SM formation. We have identi fied a number of Atg proteins that influence the trafficking of Atg9. This study reveals functiona l motifs that are essential for to pexophagy. These motifs appear critical to At g9 trafficking and function. Furt hermore, to better understand the diverse role of Atg9, its involvement in autophagy and pexophagy has been explored. The roles of a number of Sec proteins in au tophagy have been reported. These proteins are normally required for vesicular movements between the endoplasmic reticulum and Golgi apparatus in autophagy. The data have shown that ScAtg8 interacts with three v-SNAREs (Legesse-Miller et al. 2000) and eight Sec proteins are required for autophagy (Ishihara et al., 2001; Hamasaki et al. 2003; Reggiori et al. 2004). ScSec12, the guanine-nucleotide exchange factor powered by the COPII GDP-GTP exchange factor Sar1, facilitates formation of the autophagic vacuole from the PAS (Reggiori et al. 2004). ScSec16, ScSec23, and ScSec24 also function in COPII vesicle trans port of proteins and have essential roles in autophagy. The functional role of the COPII vesicle pathway in pexophagy has not been examine and forms the basis of my studies. This research highlights a role for Sar1p in pexophagy. 17

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The early membrane events leading to the onset of pexophagy are not known. The means by which the proteins involved in pexophagy are transported to their respec tive sites of activation and their activation is poorly understood. The way in which the pexophagi c proteins interact with one another is not clear. This research will elucidate the functional motifs essential to Atg9, a key transmembrane protein. These studies will al so describe the role of the COPII secretory pathway in pexophagy. Finally, this research will broach the imp lications for the maturation of the sequestering membranes whereby the pero xisomes are engulfed during micropexophagy. The Role of Atg9 in Glucose-Induced Micropexophagy The scientific researcher is resourceful. He has multiple overlapping ways to verify his hypotheses. The burgeoning field of micr opexophagy has experience basic technique development. The alcohol oxidase assay is a tool to determine whether cells undergoing glucose-induced micropexophagy are functional. Western blot allows us to study cells undergoing ethanol-induced macropexophagy. Pulse-ch ase proteolysis enables us to examine cells undergoing nitrogen starvation-induced macroautophagy. Using these methods, we have learned that Atg9 is required for multiple au tophagic pathways, including glucose-induced micropexophagy, ethanol-induced macropexophagy, a nd starvation-induced autophagy. The induction of micropexophagy by glucose adaptation reveals the formation of the SM, which nucleates from sites at the vacuole. Multiple laboratory analyses have enriched our understanding of Atg9. Ultrastructural analyses of atg9 mutants during glucose-induced micropexo phagy show a block at an early event in peroxisome degradation by the vacuole: th e vacuole is apposed to the peroxisomes and slightly indents; however, the extension of the SM is not co mpleted (e.g. stunted arm-like protrusions) and the engulfment process by the va cuole stops prematurely. Indeed, examination under the electron microscope has revealed mu ltiple peroxisomes accumulate waiting to be 18

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engulfed in atg9 cells, halted as if waiting for somethi ng; control cells observed at the same timepoints exhibit the remnants of degraded peroxisomes (Chang et al., 2005). The use of GFP-tagged proteins has revealed much concerning Atg9 cellular localization in Pichia pastoris During glucose-induced micropexophagy, Atg9 trafficks from its peripheral PC9 compartment to the PVS (perivacuolar stru cture) at the base of the membranes that sequester the peroxisomes to the SM (sequester ing membranes) at the vacuole and to the PAS (pre-autophagic structure) ad jacent to the tips of the SM for formation of MIPA (micropexophagic-specific membrane apparatus). The localization of Atg9 to the PVS requires Vps15 and Atg11 (Tina Chang, unpublished observations). The expression of Atg9 changes over time. That is, the cellular levels of Atg9 increase during the first 4 hours of glucose adaptation and th en slightly decrease between 4 and 8 hours of glucose adaptation (Chang et al., 2005). Thus, Atg9 appears to be upregulated then possibly degraded during pexophagy. The degr adation or recycling of Atg9 in P. pastoris is indeterminant. Atg9 is an integral membrane protein required for an early sequestration event of pexophagy. It localizes to the SM during glucos e adaptation. Identifying amino acid domains within Atg9 required for function wi ll reveal structural motifs important to the expansion of the sequestering membranes and the role of membra ne-bound Atg9 to this process. Atg9 encodes 885 amino acids of a 102 kDa protein. The C-termi nus is asparagine-rich. PpAtg9 shares 77% identity with ScAtg9. The large central regi on containing the transmembrane domainsresidues 190-679reveal shared homology betw een the two proteins and 45% identity in that region. There are five to six transmembrane domains with the N and C termini at the cytosolic surface of the membrane (Yen et al. 2007). Some of the conserved regions of Atg9 showed up in a motif 19

PAGE 20

database search as possible functional domains (F igure 1). For example, a region within loop #3 and the C-terminal QQHKA may act as ER exit signa ls. Furthermore, there exist two putative motifs between amino acids 235-9 and 607-11 that may interact with a peroxisomal membrane protein, Pex14. Cellular Compartments of Atg9 Under normal growth conditons, PpAtg9 in P. pastoris resides in a unique peripheral compartment, PC9 (Chang et al., 2005). This compartment does not contain COXIV-GFP (mitochondria), PpSec13 (intermediate compar tment), and PpSec7 (G olgi apparatus). Furthermore, PpAtg2, PpAtg8, and PpAtg11 are absent from these vesicles. Upon glucoseinduced micropexophagy, PpAtg9 transits from the PC9 to perivacuolar st ructures (PVS) that also contain PpAtg11, but not Pp Atg2 or PpAtg8. Finally, PpAt g9 is found at the sequestering membranes (SM) that engulf the pe roxisomes (Figure 1-2). The trafficking of ScAtg9 and its organization in to discrete puncta are affected in mutant strains in which ER morphology is altered (Mar i and Reggiori, 2007), notably trafficking of ScAtg9 to the mitochondria. Peripheral ScAtg9 does not co-localize with peroxisomal and endosomal markers (Chang and Huang, 2007). The co-localization of Atg9 with the mitochondria is a matter of debate. It has been reported that only a small percentage of the protein is found bound to the mitochondria by density gradient fractionation (Mari and Reggiori, 2007); Atg9 did co-fractionate with mitochondrial marker Por1 (Yen et al. 2007). PpAtg9 does not co-localize with the mitochondria (unpublished results). Mammalian Atg9 is localized to the TGN and late endosomes; starvation causes redistributed from the TGN to more peripheral, endosomal membranes (Young et al. 2006). Studies of mAtg9 have revealed that despite its homology to th e yeast protein, its subcellular distribution is distinct. mAt g9 do not have a specialized site for autophagosome biogenesis 20

PAGE 21

(such as the PAS) but have multiple subcellu lar locations; peripheral, endosomal membranes may be the mammalian counterpart of the PAS (Young et al. 2006). The homology of proteins such as PpAtg9 is greater between P. pastoris and H. sapiens as opposed to S. cerevisiae and H. sapiens. Transport of Atg9 During Micropexophagy Requires Vps15 And Atg11 In S. cerevisiae, Atg9 localizes to the PC9 and PAS, the site of SM nucleation and expansion. In P. pastoris, Atg9 can be found at the PC9, PVS, and occasionally at the PAS. The PVS is the site of SM nucleation and expansi on, while the membrane assembly of the MIPA occurs at the PAS. Atg9 tra fficking requires multiple ATG gene products. ScAtg9 recruits ScAtg2 and ScAtg18 to the PAS (Chang and Huang, 2007). In the absence of ScAtg2 or ScAtg18, ScAtg9 movement is re stricted to the PAS (Yen et al., 2007). Anterograde movements of ScAtg9 to the PAS requires Sc Atg11, ScAtg23, ScAtg27, and actin (Yen et al., 2007). Recycling back to the PC9 requires the ScAtg1-ScAtg13 co mplex, ScAtg2, ScAtg18, and the PI3P generated by PI3K which contains ScVp s15, ScVps43, and ScAtg14 (Mari and Reggiori, 2007). In mammalian cells, the anterograde mo vements of HsAtg9 out of the TGN to the limiting membranes of the autophagosome require HsAtg1 and PI3K activity (Young et al. 2006). In P. pastoris Vps15 and Atg11 are required for Atg9 to transit from the PC9 to the PVS, while Atg2 and Atg7 are necessary for the movements of Atg9 from the PVS to the SM during micropexophagy (Chang et al. 2005). The data suggest that Vps15, Atg11, actin, Atg1, Atg23, and Atg27 have major roles in the trafficking of Atg9 into an out of the PAS/PVS. Vps15, A Component of The Phosphatidyl Inosit ol-3 Kinase (PI3K) Complex Thought To Signal The Start Of Autophagy, Is Required For Atg9 Trafficking Vps15 is a protein kinase that functions as the regulatory subunit of the PI3K complex (Kihara et al. 2001; Budovskaya et al. 2002). Vps15 is required for nucleation of the vacuole 21

PAGE 22

during micropexophagy and is involved in early expansion by affecting the trafficking of Atg9 (Stasyk et al. 1999; Chang et al., 2005). Atg9 transit to the PV S requires Vps15 during glucoseinduced micropexophagy. Atg9 and Vps15 are not thought to directly interact Instead, Vps15 is thought to act upstream of Atg9 in a regulatory fashion. Vps15 is also required for Vacuolar Protein Sorting (by definition) and for delivery of hydrolases to the lysosome-like organelle (e.g. vacuole; equivalent to the mammalian lysosome) in yeast. Deleting or mutating Vps15 can have considerable implicatio ns. A vacuolar missorting phenotype is exhibited by vps15, in which all transport of vacuolar pr oteins is inhibited, and cellular growth is slow at 28 C and arrested at 37 C (Kihara et al. 2001). Other phenotypic defects associated with vps15 include defects in osmoregulation and disturbed vacuolar segregation (Stasyk et al. 1999). Vps15 is composed of 1340 amino acids with a predicted molecular weight of 166 kDa; it runs at about 170 kDa on denaturing gels. It has four functional domains conserved amongst species important to signal transduction, protein traffi cking and protein-prot ein interactions: the myristoylation site, the kinase domain, the HEAT repeats and the WD repeats. These domains are important for the function of Vps15 and for its interaction with other proteins. The second amino acid from the N-terminus, gl ycine, is myristoylated for peripheral membrane association (Panaretou et al. 1997). Subcellular fractionation and protease protection assays indicate that ScVps15p is associated with the cytoplasmic fa ce of an intracellular membrane. ScVps15 is myristoylated in vivo (Herman et al., 1991a; Herman et al., 1991b). Non-myristoylated Vps15 protei ns are 35-50% phosphorylated in vivo compared to the wild-type protein and possess near wild-type levels of biological activity. Non-myristoylated Vps15 still seems to be biologically and exhibi ts near wild-type growth rates. 22

PAGE 23

Amino acids 147-152 (DIKSEN) in ScVps15 is the six amino acid sequence characteristic to the serine/thre onine (S/T) kinase family; the lysine residue is absolutely conserved in S/T kinases. D147 and D165 are the aspartate residues t hought to bind the ATP phosphate groups through an intermediate Mg2+ ion. The glutamate residue occurring at residue 199 is highly conserved among protein kinases. Its mutation abolishes ScVps15s autophosphorylation (Kihara et al. 2001) and confers temperature sensitivity and causes carboxypeptidase Y missorting (Bryant and Stevens, 1998). HEAT repeats (for H untington, e longation factor 3, a lpha regulatory subunit of protein phosphatase 2A, and T OR1) form pairs of -antiparallel rod-like (hai rpin) helices and function as protein-protein interacting surfaces. The HEAT repeat is a tandemly repeated module occuring in cytoplasmic proteins involved in intracellular transport pr ocesses. Structural analyses reveal that the motif could provide a platform for prot ein-protein interactions. The HEAT repeat motif occurs three times in Vps15, between amino acids 424-462, 562-600, and 601-639. Mutations that disrupt the integrity of the HEAT repeats result in the inactivation of ScVps15 (Budovskaya et al. 2002). The last 400 amino acids of Vps15 are within the WD repeat region. WD motifs have a conserved core of approximately 40 amino acids with a GH dipeptide 11-24 residues from the Cterminus and a WD dipeptide at the C-terminus ; thus, WD40 specifies the motif type and the number of residues present in this motif. All WD-repeat proteins are thought to form circularized -propeller structures. The -propellers serve as special ized docking platforms for other proteins and require at l east four repeats for functional relevance. Two or three motifs present in single polypeptides may dimerize to form a fouror six-bladed propeller. 23

PAGE 24

ScVps15 C 214 was not phosphorylated in vivo suggesting that these amino acids contain the site of phosphorylation; deletions of this domain resu lt in a temperature-conditional vacuolar protein sorting defect which is reversed when mutant cells are shifted back to a permissive temperature. Deleti ng a short portion of the C-termi nus in addition to mutating the kinase domain results in a more se vere sorting phenotype; in addition, ScVps15ts alleles have equally severe CPY sorting def ects, and if the C-terminus is truncated, an even greater exaggeration of the sorting defect re sults. It is unclear whether the C 214 phenotype is due solely to partial deletion of the WD40 domain, or if the last 214 amino acids harbor amino acids of particular importance. ScVps15p, the regulatory subunit for the PI3K complex, and ScVps34p, the catalytic subunit for the PI3K complex, interact. Vps34 is known to be a major player in signal transduction. The gene product of VPS34 is re quired for endocytosis and for sorting to the vacuole (Stasyk et al., 1999) in Saccharomyces cerevisiae The phosphatidyl inositol (PtdIns) 3kinase (PI3K) activity of ScVps34p must be activ ated by the kinase activity of ScVps15p (Stasyk et al. 1999). An intact Vps15p prot ein kinase domain is necessary for the association with and activation of Vps34p (Stack et al. 1995). Yeast Two-Hybrid ma pping experiments identified all three HEAT repeats to be required for binding of ScVps15 to ScVps34 (Vps15 fusions retaining one HEAT repeat exhibited in termediary binding) (Budovskaya et al., 2002). The HEAT repeats are required but not sufficient fo r binding of Vps15 to Vps34. Once ScVps34p is activated by ScVps15, PtdIns molecules in the lipid bilayer (of an as yet unid entified intracellular membrane) are phosphorylated; localized patche s of these PtdIns(3)P then activ ate effector molecules, which may include Atg18, Atg20, Atg21, Atg24 or Gsa13, th at bind to these glycolipids (Stasyk et al. 24

PAGE 25

1999). The interactions of membrane-bound Sc Vps15p and ScVps34p are critical to the targeting of acid hydrolases to the yeast vacuole. Identifying the localization of Vps15 and determining the motifs required for this localization are basic to elucidating the role of Vps15 in early pexophagy. Its interactions with other proteins and its potential as a gatekeep er to pexophagic sequestration will help explain autophagy in systems besides just methylotropic yeasts. Atg11 Is a Membrane-Associated Protein Required For Pexophagy But Not Autophagy Atg11 has been shown to interact directly wi th Atg9, although the exact mechanism by which these interactions occur and the implications of these interactions may not be fully realized. Different avenues of investigation of Atg9 and Atg11 have revealed the regions necessary for these proteins to interact with one another. Immunofluorescent studies in S. cerevisiae have shown that ScAtg9 and ScAtg11 colocalize. ScAtg9 and ScAtg11 do not co-fractiona te with any other known resident organelle marker proteins (Kim et al., 2002). It has been suggested that the presence of ScAtg11 at the PAS recruits additional ScAtg9 to the same place (Kim et al., 2002). Studies in S. cerevisiae provide evidence that Atg11 inter acts with other proteins (namely Atg9) as a gatekeeper for sequestration by the vacuole. Atg9 and At g11 are known to be present at the PVS in P. pastoris. Atg9 requires Atg11 to localize to the PVS. Thus, PpAtg11 may regulate protein trafficking to the PVS. Such transport may be mediated by actin, since Reggiori and coworkers have shown that ScAtg11 interacts with actin and both are required for anter ograde transport of ScAtg9 from the PC9 to the PAS during autophagy (Monastyrska et al. 2006). Even though the localization of ScAtg9 is de pendent on ScAtg11, the reverse is not true. Localization of GFP-ScAtg11 to the PA S is not significantly affected in Scatg9 cells (Chang 25

PAGE 26

and Huang, 2007). Thus, ScAtg11 appears to a ffect PAS function independently of its interaction with ScAtg9 (Chang and Huang, 2007). PpAtg11 is a peripheral memb rane protein that contains 1313 amino acids and has a predicted molecular weight of 151,077 Daltons. Th e primary putative structural motif in Atg11 is its coiled-coil domain. The coiled-coil domain is a scaffold for prot ein interactions. Atg11 contains four coiled-coil domains (Chang and Huang, 2007), of which the first 2 coiled-coil regions of ScAtg11 are required for interaction with ScAtg9 (C hang and Huang, 2007). Deletion of residues 272-325 of Atg11 CC1 was important to Atg9-Atg11 interactions (Chang and Huang, 2007), contrary to Klionsky s earlier findings regarding th e deletion of residues 272-321. The interaction of ScAtg11 with ScAtg9 via the coiled-coil domains is essential for the recruitment of ScAtg9 to the PAS by ScAtg11. The region within Atg9 that is required for its interaction with Atg11 has been narrowed down as well. Deletion of the fi rst 152 residues of the N-terminus did not affect the interaction of Atg9 with Atg11 (Chang and Huang, 2007). De letion of the first 201 residues of the Nterminus of ScAtg9, however, did obliterate it s interaction with Atg11. (Chang and Huang, 2007). Two hybrid analyses narrowed this regi on to amino acids 154-200. Based on the fact that Atg9 154N and Atg9 201N both migrated faster than Atg9 200C, there appears to a posttranslational protein modification at the N-terminus of Atg9 which inhibits the migration of Atg9 even when the last 200 amino acids of At g9 are deleted (Chang and Huang, 2007). The Atg11 coiled-coil domains may not be the on ly regions that interact with Atg9. The transport of Atg9 after knocki ng out Atg1 (TAKA assay) in S. cerevisiae examines the localization of Atg9 to the PAS in mutant Atg1 background as control. A double knockout background determines whether Atg9 localization changes when another protein in addition to 26

PAGE 27

Atg1 is knocked out. If Atg9 is restricted to th e PAS, the gene product th at is knocked out in addition to Atg1 acts after Atg1 in Atg9 trafficking. If Atg9 does not loca lize to the PAS and is present in cytosolic dots, the ge ne product that is knocked out in addition to Atg1 acts prior to Atg1 in Atg9 trafficking. The TAKA assay has been applied to determin e whether the coiled-coil domains are the means by which Atg11 interacts with Atg9 prior to Atg1 involvement. Atg11 CC1 or Atg11 CC2 did not restore the GFP-Atg9 restri ction to the PAS during starvation in atg1 atg11 double knockout cells (Chang and Huang, 2007). Thus, it appears th at the coiled-coil domains of Atg11 are not the only region that interact with Atg9. Deletion and insertion mutations of the PpAt g11 gene product have enabled researchers to pinpoint the step at which PpAtg11 participates in micropexophagy. Both Ppatg11 (identified by the REMI mutant approach (Schroder et al. 2007)) and Ppatg11 exhibit defects in micropexophagy at an early stage of the engulfm ent process, characte rized by the vacuolar extensions only partially surrounding the pe roxisomeperhaps a checkpoint stop in the sequestration process (Kim et al., 2001). PpAtg11 also requires PpVac8 for its proper localization; Ppvac8 exhibits a completely round vacuol e characteristic of a very early micropexophagy mutant. Atg11 has a specific role in autophagy Atg11 is the only protein known to function exclusively in micropexophagy in Pichia pastoris (Kim et al., 2001). ScAtg11, which shares significant coiled-coil domain sequence homolog y to PpAtg11, is specific to the pexophagy pathway but not the macroautophagy pathway (Kim et al., 2001). PpAtg11 has an important role in pexophagy. It distributes around the peroxisomes early in sequestration, possibly tagging them fo r vacuolar recognition during pexophagy. Upon 27

PAGE 28

induction of pexophagy, localization of PpAtg11 to the point of contact between the vacuolar and the peroxisomal membranes occurs (Kim et al., 2001). PpAtg11 is concentrated in the PVS adjacent to the vacuole membrane, and it has be en hypothesized that this protein mediates interactions between the vacuole and the peroxisome during microautophagy (Kim et al., 2001). Based on the existence of multiple bands by We stern blot, PpAtg9 is thought at least to dimerize (unpublished observations). Atg11, as well, is thought to dimerize, at least in S. cerevisiae. ScAtg11p forms detergent -resistant homodimeric complexes (or homooligomers) (Kim et al., 2001). The presence of multimeric Atg11 complexes has not been confirmed in P. pastoris, but their formation via the coiled-coil domain may be important to PpAtg11s association with itself and other proteins during micropexophagy. Another motif present in PpAtg11 that may be important to its functi on is the WKSYY sequen ce (amino acids 241-5), a WXXXF/Y motif shared by Pex5, Pex14 and At g9. This domain may enable PpAtg11 to interact with Pex and other Atg proteins. Once Pex14 or another protein attracted to the PpAtg11 WXXXF/Y motif, the coil ed-coil domain of PpAtg11 may promote and enhance this interaction, possibly resulting in multimeric complex formation. Actin Is Required For Efficient Autophagy It is possible that Atg11 dire cts the trafficking of Atg9 to the PVS via actin cables (Monastyrska et al., 2006). A role for actin has been implicated in autophagy. Atg9 and Atg11 trafficking is altered and the formation of the PA S is aberrant when toxins or mutant strains inhibit actin polymerization (Reggiori et al., 2003; Reggiori and K lionsky, 2006; Mari and Reggiori, 2007; Yen et al. 2007). Lantrunculin A destroys th e actin cytoskeleton by interfering with polymerization. ScAtg9 does not accumulate at the PAS in atg1ts strain treated with lantrunculin A (Yen et al. 2007). In the absence of Atg11 and actin filaments, ScAtg9 28

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anterograde transport from the PC9 to the PAS is blocked [Mari Reggiori Atg9 trafficking in S. cerevisiae Autophagy 2007]. However, Atg11 an d actin filaments are not required for Atg9 transport to the PAS during starvation-induced autophagy (Mari and Reggiori, 2007). Thus, actin is required for Atg9 movement duri ng normal growth and pexophagy but not starvationinduced autophagy. Moreover, a toxin or a mutant strain that blocks actin filament formation severely impairs Atg9 transport from the PC9 locations (Mari and Reggiori, 2007), suggesting that the initial activation of Atg9 is impaired. Another requirement for actin has been described in regards to Atg11. ScAtg11 is mislocalized from the PAS to several disperse d fluorescent dots throughout the cytoplasm in VFT (Vps53, V PS F iftyt hree) mutants (Reggiori et al. 2003). The VFT complex, named for Vps53, the first protein discovered in the complex, functions as a tetherin g factor for retrograde traffic from the early endosome back to the late Golgi and is required speci fically for degradation processes during normal growth in S. cerevisiae. Nitrogen starvation-i nduced autophagy elicits the correct re-localization of ScAtg11and the block in VFT mutants is bypassed. Further investigation showed that Vps51, Vps52, Vps53 and Vps54 are required for actin organization (Reggiori and Klionsky, 2006). The Atg1 Protein Kinase Is Requi red For Pexophagy And Autophagy Atg1 is required for late expansion during micropexophagy. Atg9 localization late in autophagy requires Atg1 in S. cerevisiae. Whether Atg1 acts after Atg11 is no longer a matter of debate. GFP-Atg9 is restricted to th e PAS in growing or starved cells in atg1 cells (Chang and Huang, 2007). The transport of Atg9 after knocking out Atg1 (TAKA assay) in S. cerevisiae examines the localization of Atg9 to the PAS in mutant Atg1 background as control. 29

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Deletion of full-length Atg11 and Atg1 together provide further in sight regarding Atg9 trafficking. In atg1-atg11 background, GFP-Atg9 localized to the PC9, not the PAS. Similar results were obtained in atg1-atg23 and atg1-atg27 cells. For example, in atg27 and atg27-atg1 cells, Atg9 localized to multiple punctate dotsnone of which corresponded to the PASand some of which were re stricted to the mitochondria (Yen et al. 2007). Atg27 may be required for movement of Atg9 from the mito chondria to the PAS and is thought to function before Atg1 in the cycling of Atg9 (Yen et al., 2007). Thus, different null Atg proteins were substituted and the atg1 deletion was constant during the TAKA assay, and the action of Atg23 and Atg27 in regards to Atg9 tra fficking was determined. Taken together, this would suggest that Atg11, Atg23, and Atg27 are required for anter ograde transport from the PC9 to the PAS, and that Atg1 is the only protein known to be required for retrograde transport The majority of Atg27 is restricted to the PAS in atg1 cells (Yen et al., 2007). In atg9 and atg1 -atg9 cells, Atg27 localized to multiple punctate structures similar to wild-type localization except that none of the dots localized to the PAS. In this case, Atg9 and Atg1 were deleted and the localization of Atg27 to the PA S was studieda modified TAKA assay with an aim to determine Atg27 trafficking. Atg9 is required for anter ograde movement of Atg27 to the PAS (Yen et al., 2007). Retrograde movement of Atg27 to the PC9 is dependent on the Atg1Atg13 complex (Yen et al. 2007), however Atg1 kinase activity was not required. The region for Atg27 and Atg9 interactions is being whittle d down: the anterograde movement of Atg9 to the PAS does not require the cytosolic C-terminus of Atg27 (Yen et al., 2007). Atg9 dots co-localize with Atg8 and At g5, which are known to be involved in autophagosome nucleation. Thus, the multiple G FP-Atg9 dots induced by starvation treatment of atg1-ape1 cells is thought to repres ent nucleated sites of au tophagosome assembly (Chang 30

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and Huang, 2007). GFP-atg9 154-201 behaved the same as GFP-Atg9 during growth or starvation in atg1-ape1 background (Chang and Huang, 2007). Atg9 Retrograde Trafficking Requires Atg23 And Atg27 Atg27, Atg23 and Atg11 are essential fo r Atg9 retrograde trafficking (Yen et al., 2007). Atg9 and Atg23 show similar distribution to Atg27 lo calization, with non-PAS cytosolic dots. Atg23 is a cytosolic protein required for the Cvt pathway and autophagy but not pexophagy (Yen et al., 2007). Because there is no e quivalent Cvt pathway in P. pastoris and the role of Atg23 in P. pastoris is unclear, for our purposes Atg23 is negl ected. PpAtg27 is an integral membrane protein required for efficient pexophagy, which ma y be related to its Atg9 trafficking role. In addition, Atg27 is required fo r starvation-induced autophagy. atg27 cells exhibited partial induction of Pho8 60 activity (a biochemical measur e of normal autophagy), suggesting that starvation-induced autophagy did occur but not as efficiently as in wild-type cells (Yen et al. 2007). With further investigation, atg27 was shown to have reduced autophagosome number (a defect in nucleation) or reduced au tophagosome size (a defect in expansion) (Yen et al. 2007). Thus, Atg27 is required for nucle ation and expansion during macroautophagy. Structure precludes function; t hus, the structure of Atg27 is described in detail. Atg27 (also known as Etf1) is a Type I transmembrane protein (Yen et al. 2007). Atg27 topology indicates that it could be present within th e intermembrane space betw een the autophagosome inner and outer vesicle membrane or in the lu menal space during autophagosome formation (Yen et al. 2007). The C-terminus of Atg27 is cytosolic (Yen et al. 2007). Atg27 harbors an Nterminal signal sequence and the N-terminus may therefore also face the cytoplasm (Yen et al., 2007). 31

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Atg27 cellular localization has been examin ed fluorescently. Atg27GFP distributes to several subcellular punctate stru ctures. None of these was found to co-localize with the ER or with the peroxisomes (Yen et al., 2007). ScAtg27 does co-locali ze with the PAS marker Ape1 (Yen et al., 2007), with MitoFluor red (which labels the mitoc hondria) and with the Golgi marker Vrg4-GFP (Yen et al., 2007). Atg27 cellular localization is similar to Atg9 localization except that Atg9 has not been observed at the Golgi. To verify the fluorescence data, Atg27 vesi cles were separate d by sucrose density gradient fractionation (Yen et al., 2007) and co-fractionated with Por1 and the Mnn1 marker for the Golgi complex (Yen et al., 2007). Multiple membrane sources are thought to supply the lipids required for autophagosome formation; these lipids may orig inate in the Golgi. Atg27 is a putative PI3P binding protein downstream of Vps34 (Yen et al. 2007). The Vps34 kinase complex I generates PtdIns(3)P at the PAS and is essential for autophagy (Yen et al. 2007), specifically MIPA formation. T hus, Atg27 may regulate lipids esse ntial to MIPA formation. Atg27 is required for Atg9 cycling between the PAS, mitochondria and Golgi (Yen et al. 2007). The localization and tr ansit pattern of Atg27 is similar to that of Atg9 (Yen et al. 2007). Atg9 was predicted to interact with Atg27 by y east two hybrid analysis. Affinity isolation showed that Atg9 and Atg27 inte ract with one another (Yen et al. 2007). Components Of The COPII Secretory Pathway Are Required For Autophagy A number of Sec proteins ha ve been shown to be require d for macroautophagy (Table 12). Notably, Klionsky showed that Sec12, whic h is the guanine nucleo tide exchange factor (GEF) for Sar1p recruiting this protein to the ER membrane, is required for the formation of the autophagic vacuole (Ishihara et al. 2001; Reggiori et al. 2004). Sec18 and Vti1 are required for fusion of the autophagic vacuole with the yeast vacuole. Finally, Sec7, Sec17, and three COPII vesicle coat proteins (Sec16, Sec23, and Sec24) are necessary for autophagy, but their 32

PAGE 33

roles have not been defined (Ishihara et al., 2001; Hamasaki et al., 2003). Sec12, Sar1, Sec23 and Sec24 are required for assembling the cargo to be packaged by the COPII vesicle (Sato and Nakano, 2007). Interestingly, Sec13 and Sec31 esse ntial for polymerization of the COPII coat were not required for autophagy (Ishihara et al. 2001). The data suggest that components of COPII assembly, but not COPII vesicles, are e ssential for autophagy. However, the roles of COPII vesicles and of Sar1, a major player in the assembly of the COPII vesicles, in autophagy and pexophagy have not been investigated. We have shown that that Sar1 is required for pexophagy and autophagy suggesting a direct link between the secretory pathway and these degradative pathways. Sar1 and Protein Transport COPII vesicles organize the deli very of proteins from the ER to the Golgi apparatus. Mutants that disrupt ER to Golgi transport can be categorized into two ca tegories, phenotypically defined as those defective in fusion of ER-der ived vesicles with the Golgi complex (that accumulate 50 nm vesicles but do not fuse with the Golgi) and those defective in vesicle formation (that accumulate in ER membranes). Sa r1 is the GTPase responsible for incorporating cargo into COPII vesicles. Upon accepting GTP from Sec12, active Sar1 interacts with Sec23 and Sec24 at the ER and the COPII complex a ssembles. The COPII complex assembles cargo and then polymerizes and buds from the ER. Th e hydrolysis of the GTP by the Sar1 results in COPII complex disassembly and uncoating the COP II vesicles which then fuse with the Golgi apparatus deliveri ng their contents. Sar1p-guanosine 5O -(3-thiotriphosphate), a no n-hydrolyzable form of GTP, is unable to promote targeting of ER vesicles to the Golgi at all (Oka and Nakano, 1994). In other words, GTP hydrolysis by Sar1 is not required for vesi cle formation but is required for subsequent 33

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transport from the ER to the Golgi. Cells expressing sar1(T39N) (e quivalent to T34N in P. pastoris ) has an extensive ER and ar e defective in COPII vesicle formation. Cells expressing sar1(H79G) form an exaggerated Go lgi apparatus and are defective in retrograde transport to the ER. The sar1(T39N) mutant has a high affinity for GDP and blocks COPII vesicle formation from the ER. The sar1(H79G) mutant is unable to hydrolyze GTP and prevents disassembly of COPII vesicle coats thereby inhibiting re trieval of proteins to the ER (Ward et al., 2001). Sar1 controls COPII budding and that it was hypothesized that Sa r1 is the means by which coat assembly and cargo selection are integrated (Barlowe, 2002). Membrane-bound Sar1-GTP links vesicle cargo to pre-budding comple xes and to the COPII coat. At that time it was not understood how coat polymerization leads to membrane deformation, how the variety of cargo to be included in COPII vesicles is rec ognized by coat subunits, and how Sar1 regulates the coat assembly/disassembly stages to execute these tasks (Barlowe, 2002). The recycling of proteins fr om the Golgi complex is not the principle means by which sorting is achieved: rather, th e separation of proteins destined for transport (as opposed to proteins that remain in the ER) occurs primarily at the moment of transport vesicle budding from the ER. Thus, Sar1 is critical to protein dispersion throughout the cell. Sar1 was reported in yeast to be detected in a diffuse, peri nuclar and reticular localization (overlapping with Kar2 and Sec62) as well as in punctuate struct ures that did not overlap with ER or Golgi markers (Kuge et al., 1994). These structures may be a small population of Sar1 that labels transport vesicles so that they might eventually be recycled back to the ER. Sar1 serves as a discriminating molecule that transpor ts key proteins throughout the cell and imparts a unique signal to a membrane certifying proper tr avel throughout the secr etory pathway be it to the Golgi apparatus or degradation by the ubiqu itin-proteasome system (Sato and Nakano, 2007). 34

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Thus, a role for Sar1 could be to provide regula tory control to sort a nd concentrate cargo to ensure efficient export from the ER (Kuge et al., 1994). Sar1 And Lipid Transport Ceramide is synthesized in the ER and is so rted to other organelle s in a non-vesicular fashion by at least two pathways: an ATPand cytosol-dependent major pathway and an ATPand cytosol independent minor pathway (Hanada et al. 2003). CERT, for example, interacts specifically with the ER membranes and specifically extracts ceramide. This may suggest that another lipid-transfer-catalyzing enzyme specifically extracts PtdEtn. The importance of PtdEtn trafficking is that during aut ophagy Atg8 becomes covalently bound to this lipid at the expanding autophagic vacuole. Sar1 may decrease the propen sity of PtdEtn to traffick properly to the forming autophagic vacuole. However, in cells expressing sar1(T 39N) the conversion of ceramide to sphingomyelin was not affected. This would suggest that PtdEtn trafficking would not be affected but is by no means definitive. The ATP dependence of ceramide transport could be relevant to PI4-P metabolism at the Golgi (Hanada et al. 2003). A specific subdomain of the ER is thought to be closely associated with the Golgi stacks (Hanada et al. 2003). If PtdEtn trafficks via a non-ve sicular mechanism, th e integrity and the close association of the ER and th e Golgi may be essential to PtdEtn trafficking; the alteration of this association by mutant Sar1 would alter PtdEtn trafficking or posttranslational modification of PtdEtn and conversion of PtdEtn to another compound, such as PI4-P, that is crucial for Atg8 trafficking and MIPA formation. Summation Glucose signals and peroxisome recognition in itiates the sequestration of peroxisomes by micropexophagy. During sequestration, the SM nucleat e from the vacuole, then expand from the PVS and PAS (Figure 1-1). Completion occurs when the peroxisomes are engulfed and 35

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incorporated into the vacuole where they are de graded. The sequestration of peroxisomes begins by nucleation of the sequestering membranes (S M) from the vacuole. During expansion, two unique membranes surround the peroxisomes: the SM that expands from the PVS and the MIPA that assembles from the PAS. SM expansi on from the PVS is Atg9 dependent, while the formation of the MIPA from the PAS is depende nt upon Atg8 and Sar1. The MIPA is situated between opposing membranes thereby promoti ng membrane fusion of the SM and MIPA resulting in the completion of th e sequestration of the peroxisomes. Once inside the vacuole the peroxisomes are degraded by the hyrdrolytic en zymes and the amino acids, sugars, and lipids recycled. In this thesis, I will characterize the roles of Atg9 and Sar1 in SM expansion during micropexophagy. 36

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Table 1-1: Autophagy-related Genes Gene Name Pichia pastoris2 Function3 Characteristics References ATG11 GSA10 PAZ1 P,A Serine/threonine-kinase that complexes with Atg11 and Vac8 (Stromhaug et al., 2001; Mukaiyama et al., 2002) ATG2 GSA11 PAZ7 P,A Peripheral membrane protein that interacts with Atg9 (Stromhaug et al., 2001; Mukaiyama et al., 2002) ATG3 GSA20 P,A E2-like enzyme responsible for the conjugation of Atg8 to lipids ATG4 PAZ8 P,A Cysteine proteinase (Mukaiyama et al., 2002) ATG7 GSA7 PAZ12 P,A E1-like enzyme responsible for the conjugation of Atg12 to Atg5 and Atg8 to lipids (Yuan et al., 1999; Mukaiyama et al., 2002) ATG8 PAZ2 P,A Soluble protein that becomes conjugated to lipids (Mukaiyama et al., 2004; Monastyrska et al., 2005) ATG9 GSA14 PAZ9 P,A Integral membrane protein associated with structures juxtaposed to the vacuole and essential for expansion of the sequestering membranes This Study ATG11 GSA9 PAZ6 P Coiled-coil domain found at vacuole surface (Kim et al., 2001; Klionsky et al. 2003) ATG18 GSA12 P,A WD40 protein (Guan et al., 2001) ATG24 PAZ16 P,A Sorting nexin with PX domain (Ano et al., 2005) ATG26 PAZ4 P UDP-glucose:sterol glucosyltransferase (Mukaiyama et al., 2002) ATG28 PpATG2 8 P Coiled-coil protein (Dunn et al., 2005) PFK1 GSA1 P -subunit of phosphofructokinase (Yuan et al., 1997) PEP4 GSA15 PAZ14 P,A Endopeptidase (Mukaiyama et al., 2002) VPS15 GSA19 PAZ13 P,A Membrane-anchored Serine/threonine-kinase with WD40 domains (Stromhaug et al., 2001; Mukaiyama et al., 2002) Vac8 PpVac8 P Armadillo-repeat protein (Fry et al. 2006) 1A unified gene nomenclature of autophagy and autophagy-related processes such as pexophagy has been standardized across sp ecies to be ATG for the gene name. 2 GSA, glucose-induced selective aut ophagy; PAZ, pexophagy zeocin-resistance. 3 P, required for pexophagy; A, required for autophagy; nd, not determined. 37

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Table 1-2: Secretory-related Genes Gene Name Function1 Characteristics References SEC7 A Guanine nucleotide exchange factor (GEF) for ADP ribosylation factors involved ER-to-Golgi transport (Reggiori et al. 2004) SEC12 AV formation Guanine nucleotide exchange factor (GEF) for Sar1p involved in ER-to-Golgi transport (Ishihara et al. 2001; Reggiori et al. 2004) SEC16 A formation COPII vesicle coat protein (Ishihara et al. 2001; Hamasaki et al. 2003) SEC17 A Peripheral membrane protein required for ER-to-Golgi transport (Ishihara et al. 2001) SEC18 AV fusion with vacuole N-ethylmaleimide-sensitive fusion protein, NSF (Ishihara et al. 2001) SEC23 A formation COPII vesicle coat protein (Ishihara et al. 2001; Hamasaki et al. 2003) SEC24 A COPII vesicle coat protein (Ishihara et al. 2001) SAR1 P,A GTP-binding protein of the ARF family, component of COPII coat of vesicles; required for ER-toGolgi transport This Study VTI1 AV fusion with vacuole N-ethylmalemide-sensitive fusion protein attachment protein, SNARE (Ishihara et al. 2001) 1 P, required for pexophagy; A, required for autophagy; AV, autophagic vacuole. 38

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39

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Figure 1-1. Peroxisome engulfment by glucose-induced micropexophagy. Micropexophagy occurs by specific events define d phenotypically and genetically. These events are dependent upon autophagy-related (Atg) and other protei ns. Upon glucose signaling and peroxisome recognition, the sequestering membranes (SM) nuclea te from the vacuole. The SM then expands from perivacuolar structures (PVS) and the micropexophagy appara tus (MIPA). Sequestration is complete when the peroxisomes are incor porated into the vac uole for degradation. 40

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Figure 1-2. Molecular events of the expansi on of the sequestering membranes that engulf peroxisomes during micropexophagy. There exist multip le molecular events that regulate the expansion of the sequestering membranes to engulf the peroxisomes (P). Upon glucose adaptation, nucleation of the seque stering membranes proceeds from the vacuole (V). We have subdivided the expansion events into two categories: early Atg9 dependent and late Atg8 dependent. The early events involve the associat ion of Atg2 with a uniqu e sorting compartment (SC), which requires Atg9, Atg18 and Vps15. At this time, Atg9 transits from a peripheral compartment (PC9) to perivacuolar structures (PVS) and the sequestering membranes (SM). The trafficking of Atg9 requires Vps15, Atg11, Atg2 and Atg7. During late expansion, the SM are tethered for fusion by the micropexophagy apparatus (MIPA). The assembly of the MIPA from the pre-autophagosomal structure (PAS) requi res Atg8. The membrane anchoring of Atg8 by lipidation is mediated by At g3, Atg4, and Atg7, which in turn is regulated by Vps15. Upon fusion of the SM with the MIPA, the peroxisomes are incorporated into the vacuole where they are degraded. 41

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CHAPTER 2 IDENTIFICATION OF PEXOPHAGY GENES BY RESTRICTION ENZYME-MEDIATED INTEGRATION (REMI)1 Introduction The methylotropic yeast, Pichia pastoris can grow on methanol by synthesizing peroxisomal (e.g., alcohol oxidase and dihydroxy acetone synthase) and cytosolic (e.g., formate dehydrogenase) enzymes necessary to metabolize and assimilate this carbon source. Upon adapting these cells from growth on methanol to a medium containing glucose, the peroxisomes are rapidly and selectively degraded by a process called micropexophagy (Sakai et al. 1998; Stromhaug et al., 2001; Mukaiyama et al., 2002; Habibzadegah-Tar i and Dunn, 2003). During glucose adaptation, clusters of peroxisomes are surrounded by arm-like extensions from the vacuole. Upon homotypic membrane fusion of the arms, the peroxisome cluster is incorporated into the vacuolar lumen where it is subsequently degraded. The highly regulated degradation of these la rge peroxisomes combined with classical yeast genetics makes P. pastoris an ideal model to characterize the molecular events of pexophagy. In order to identify those genes es sential for pexophagy, we developed a direct colony assay by which to screen thousands of nitrosoguanidine-generated mutants that were unable to degrade peroxisomal alcohol oxidase Once the mutant was verified, we would identify the mutated gene by complementation with a genomic library. This approach was quite time consuming and false positives were inevitabl e. Therefore, we set out to design a new approach to identify those genes es sential for glucose-induced pexophagy. 1 Article reprinted with permission from publisher: Schroder, L.A., B.S. Glick, and W.A. Dunn, Jr. (2007) Identification of pexophagy genes by restriction enzyme -mediated integration (REMI). In Methods in Molecular Biology (Vol 389): Pichia Protocols, Second Edition. J.M. Cregg, editor. Humana Press. pp. 203-218 42

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We have utilized restriction enzyme-mediate d integration (REMI) to facilitate a random disruption of genes in the P. pastoris genome by the incorporation of a Zeocin resistance gene. P. pastoris was transformed with a pREM I-Z vector that had been lin earized with BamHI in the presence of BamHI or DpnII that would ra ndomly cleave genomic DNA leaving four base overhangs compatible with the linearized vector. Transformed cells were selected by growth on Zeocin plates. Those transformed cells unable to degrade peroxisomes during glucose adaptation were identified by direct colony a ssays and the interrupted genes ( GSA, glucose-induced selective autophagy) sequenced. REMI mutagenesis provides a novel ap proach to identifying key genes involved in autophagic pr ocesses and straightforward assays estimate the impact of the interrupted gene on pexophagy. Materials Pichia Pastoris and E. coli Strains 1. GS115 ( his4 ) 2. PPF1 ( his4 arg4 ) 3. DH5 Culture Media 1. YPD: 1% Bacto yeast extract, 2% Bacto peptone, and 2% dextrose. 2. YND: 0.67 % yeast nitrogen base, 0.4 mg/l biotin, and 2% glucose. 3. YNDH: 0.67 % yeast nitrogen base, 0.4 mg/l biotin, 40 g/l histidine, and 2% glucose. 4. YPD+Z: 1% Bacto yeast ex tract, 2% Bacto peptone, and 2% dextrose plus 100 g/ml Zeocin. 5. YNM: 0.67 % yeast nitrogen base, 0.4 mg/l biotin, and 0.5% methanol. 6. YNMH: 0.67 % yeast nitrogen base, 0.4 mg/l biotin, 0.5% methanol, and 40 g/l histidine. 7. LB+Z: 0.5% Bacto yeast extract, 1% B acto tryptone, and 0.5% NaCl plus 25 g/ml Zeocin. 43

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8. YETM: 0.5% Bacto yeast extract, 1% Bacto tryptone, and 5% MgSO47H2O, pH to 7.5 with KOH. Vector 1. pREMI-Z (NCBI accession number AF282723) Transformation of Pichia Pastoris 1. YPD 2. 1M NaHEPES, pH 8 3. 1M dithiothreitol 4. 1M Sorbitol Qualitative Assessment of Alcohol Oxidase Degradation by Direct Colony Assay 1. P5 Filter Paper, 9 cm circle 2. NitroBind, nitrocellulose 0.45 m, 85mm circle 3. AOX Detection Solution: 3.4 U/ mL horseradish peroxidase 0.56 mg/mL 2,2-azinobis (3ethylbenzthazoline-6-sulfonic acid), 33 mM potassium phosphate buffer pH 7.5, 0.13% MeOH. 4. Liquid Nitrogen Quantitative Assessment of AOX De gradation by Liquid Medium Assay 1. Cell Lysis Buffer: 20 mM Tris pH 7.5, 50 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 1 g/mL pepstatin A, 0.5 ug/mL leupeptin. 2. 425-600 m glass beads (Sigma G-8772) 3. AOX Assay Solution: 3.4 U/mL horseradish peroxidase, 0.56 mg/mL 2,2-azinobis (3ethylbenzthazoline-6-sulfonic acid), 33 mM potassium phosphate buffer pH 7.5. 4. MeOH 44

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5. 4N HCl Isolation of Yeast Genomic DNA 1. DNA Extraction Buffer :10 mM Tris pH 8.0, 2% Triton X-100, 1% SDS, 100 mM NaCl, 1 mM Na2EDTA. 2. Phenol:chloroform:isoamyl alcohol (25:24:1 v/v) 3. Tris EDTA (10 mM Tris pH 7.5, 1 mM Na2EDTA) 4. 10 mg/ml RNase A 5. Chloroform:isoamyl alcohol (24:1 v/v) 6. 3M NaOAc 7. 425-600 m glass beads (Sigma G-8772) 8. Ethanol (ice cold) Methods We have developed a protocol to rapidly identify genes e ssential for glucose-induced micropexophagy (see Figure 2-1). This method utilizes a random integration assisted by restriction enzymes of a Zeocin resistance cassette vector into the genomic DNA thereby disrupting gene expression. The Zeocin-resistant yeast defective in peroxisome degradation are then identified by direct colony a ssays. Afterwards, the site of insertion of the REMI-Z is determined by first digesting the genomic DNA with restriction enzymes and then isolating the pREMI-Z with flanking genomic DNA by amplifying in E. coli Once this vector is isolated, the flanking genomic DNA can be sequenced and the gene identified by BLAST searches of the NCBI databases. The methods described below outline: 1. vector construction, 2. yeast transformation with the pREMI-Z vector, 3. identification and isol ation of pexophagy mutants, 4. isolation of the pREMI-Z vector with flanking genomic DNA, and 5. identification of mutated genes. 45

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1.0. Vector Construction A pREMI-Z vector containing the Ze ocin-resistance gene behind the S. cerevisiae TEF and bacterial EM7 promoters to facilitate expression in yeast and bacteria (E. coli ) was constructed. This includes (a) construction of vect or and (b) amplification of vector. Two 68 base oligos were annealed to yield th e adapter that contains multiple stop codons in all 6 frames and the M13 reverse and M13 (-20) sequencing primers pointed in towards the central BamHI site: GATCGGAAACAGCTATGACCATGTCAGTCAGTCA GGATCC TAGCTAGCTAGACTGGCCGTCGTTTTAC CCTTTGTCGATACTGGTACAGTCAGTCAGT CCTAGG ATCGATCGATCTGACCGGCAGCAAAATGCTAG The parent vector pPICZ-A (Invitrog en, San Diego) was cut at unique BamHI and BglII sites. The above adapter was then lig ated into the pPICZ-A fragment at the BamHI and BglII sites to create pREMI-Z (see Figure 2-1). The vector was amplified by transforming Rbcompetent DH5 and growing the transformants on LB+Z. 2.0 Yeast Transformation The next step in this process involves the transformation of yeast and genomic integration of the pREMI-Z vector. This se ction includes (a) prep aration of electro-com petent yeast, (b) transformation yeast by electroporat ion, (c) selection on minimal me dium and (d) verification of vector integration. 2.1 Electro-Competent Yeast 1. GS115 of PPF1 cells are grown in 250mls YPD to an OD600 of 1.0 1.5. 2. Cells are harvested at 4 C in sterile 250 mL Nalgene centrifuge bottles at 4000xg for 20 minutes. 46

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3. Cells are then resuspended in 50mls YPD containing 1ml 1M HEPES pH 8. 1.25 mls of 1M DTT is added dropwise while swirling th e cells and the cells are incubated at 30 C for 15 minutes with gentle shaking. 4. Bring to 250mls with sterile cold wa ter and harvest the cells as above. 5. Wash cells two times with 250mls of sterile cold water. 6. Resuspend cells in 25mls sterile cold 1M sorbitol, transfer to 50 ml conical tube and harvest cells at 2000xg for 10 minutes. 7. Resuspend cells into 0.5mls sterile cold 1M sorbitol. Electro-competent P. pastoris are more effective when used imme diately, but can be frozen at -80 C for later use. 2.2 Transformation of Yeast by Electroporation 1. pREMI-Z is linearized by digestion with BamHI. 2. 50 L of competent yeast cells are mixed with 1-2 g of linearized pREMI-Z and 0.5-1.0 units of BamHI or DpnII. 3. The solution is then transf erred to a 0.2 cm gap cuvette for electroporation (1.5 kV, 25 F, 400 Ohms). 4. Immediately after electroporati on the cells are suspended in 1 mL of cold 1M sorbitol. 2.3 Selection of Transformants 1. The entire sample of electroporated cells is transferred to multiple YPD+Z plates (200250 L per plate). 2. Plates are then incuba ted for 2-4 days at 30 C until colonies appear. 2.4 Verifying Vector Integration 1. Pexophagy mutants are grown to stati onary growth in 2 mL of YPD 47

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2. Cells are lysed and genomic DNA isolated as described in Section 3.4.1 3. 10 L of genomic DNA is digested with EcoRI 4. The genomic DNA fragments are then separate d on an agarose gel and transferred to nylon membranes. 5. The DNA probe was made from BamHI-linea rized pREMI-Z following instructions included in the North2South Biotin Random Primer Kit (Pierce, Rockford, IL) 6. pREMI-Z vectors containing DNA fragments on the nylon membranes were detected following instruction included in the North2South Chemiluminescent Hybridization and Detection Kit (Pierce, Rockford, IL). 7. The visualized bands demonstrate pREMI-Z in sertions (Fig. 2-2). Virtually all the Zeocin-resistant clones have a single band suggesting a single insertion site. However, it appears that some clones have pREMI-Z insert ed into two genes (see R23, Fig. 2-2). 3.0. Identification and Isolation of Glucose-induced Pexophagy Mutants After isolating hundreds to thousands of Zeocin-res istant clones, the next step is to screen for pexophagy mutants. This is done by direct co lony assay to identify those clones that do not degrade AOX when adapted from methanol to gl ucose. Those pexophagy mutants identified by direct colony assay are then isolated and ve rified by a liquid medium assay. This section includes (a) identificatio n of pexophagy mutants by direct col ony assay and (b) quantification of the pexophagy defect by liquid medium assay. 3.1 Direct Colony Assay 1. Colonies are grown on YPD+Z selecti on plates overnight 2-3 days at 30 C. 2. Colonies are replica-plated from the YPD+ Z plates to YNM plates (with appropriate amino acid supplements) and grown for 3-4 days at 30 C. 48

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3. Colonies are then replica-plat ed from the YNM plates to ni trocellulose which is placed onto YND plates colonies up and grown for 16-18 hours at 30 C. 4. The colonies on nitrocellulose are removed from the YND plat es and the cells lysed by freezing in liquid nitrogen for 20 seconds, taking care not to break the paper into shards. 5. The frozen nitrocellulose is carefully placed on Whatman paper soaked with AOX Detection Solution (33 mM potassium phos phate buffer (pH 7.5) with 0.13% MeOH, 3.4 U/mL HRP, and 0.53 mg/mL ABTS). 6. The nitrocellulose is incubated for 60 -90 minutes at room temperature. 7. AOX activity is visualized with the appearance of periwinkle (purple) dots (Figure 2-3). When grown on YNM plates, all colonies have AOX activity. However, only those pexophagy mutants unable to degrade AOX will have AOX activity when transferred from YNM to YND. 3.2 AOX Degradation by Liquid Medium Assay 1. Pexophagy mutants identified by direct colony assa y are isolated either from the original YPD-Z or the YNM plates and grown in YPD medium. 2. Cells (0.6 mL of saturated YPD culture) ar e grown in 20 mL YNM with appropriate amino acid supplements. 3. After 36-38 hours growth on YNM, 0.4 g glucose is added. 4. Two mL aliquots of cells at 0 and 6 hour s of glucose adaptation are harvested by centrifugation. 5. The pellets are resuspended in 1 mL of cell lysis buffer (20 mM Tris pH 7.5, containing 50 mM NaCl, 1 mM EDTA, 1 mM PMSF, 1 g/mL pepstatin A, and 0.5 g/mL leupeptin). 49

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6. The cells are then lysed by vortexing in the presence of 0.5 g of 425-600 m glass beads. 7. The glass beads and cellular de bris are removed by centrifugation. 8. Alcohol oxidase is measured by adding 50 L of this extract to 3 mL of AOX Assay Solution (3.4 U/mL HRP and 0.53 mg/mL ABTS in 33 mM potassium phosphate buffer, pH 7.5). 9. The reaction is started by adding 10 L MeOH and continued at room temperature for 15-30 minutes until the appearance of a teal color. 10. The reaction is stopped by addition of 200 L 4N HCl. 11. The developed teal color is measured at 410 nm. A representation of AOX degrada tion measured by liquid medium can be seen in Figure 2-4. Parental GS115 cells degrade over 90% of th e AOX after six hours of glucose adaptation compared to 10% in gsa7 and pep4/prb1 mutants. In comparison, 30-60% of the AOX is degraded by eight unique REMI mutants. 4.0 Identification of the Disrupted Gene caused by the Insertion of pREMI-Z After isolating the pexophagy mutants and ve rifying on Southerns (see Section 3.2.4) that the pREMI-Z inserted into a single site, the next step is to identify the site of insertion and the disrupted GSA/PAX gene. This is done by isolating the genomic DNA, digesting the DNA with selected restriction enzymes, ligating the pREMI-Z and its fla nking DNA into a circular vector, and amplifying the vector in E. coli for DNA sequencing. This section will cover (a) the isolation of gDNA in yeast cells, (b) the isolation of the pREMI-Z vector, and (c) sequencing the genomic DNA that flanks the pREMI-Z. 4.1 Genomic DNA Isolation 1. P. pastoris is grown to stationary phase in 1 mL YPD at 30 C. 50

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2. Harvest the cells by centrif ugation (14,000xg; 2 minutes). 3. To the cell pellets add 0.2 mL of DNA extraction buffer (2% Triton X-100, 1% SDS, 100 mM NaCl, 10 mM Tris-HCl pH 8, and 1 mM Na2EDTA), 0.2 mL of phenol:chloroform:isoamyl alc ohol (25:24:1), and 0.3 g 425-600 m glass beads. 4. Vortex on high setting for 3-4 minutes. 5. Add 0.2 mL of Tris-EDTA pH 8. 6. Microfuge (14,000xg; 2 minutes). 7. Transfer the top, aqueous layer to a fresh 1.5 mL microfuge tube and add a 2:1 volume of 100% ethanol (about 1 mL) and gently mix by inversion. 8. Microfuge (14,000xg; 2 minutes) and decant supernatant. 9. Resuspend the pellet in 0.4 mL Tris-EDTA pH 8 and 3 L 10 mg/mL RNase A. 10. Incubate 5 minutes at 37 C. 11. Add 10 L 4M ammonium acetate and 1 mL 100% cold EtOH and gently mix by inversion. 12. Microfuge (14,000xg; 2 minutes) and decant supernatant. 13. Air-dry the pellet (approximate ly 20 minutes; do not allow the pellet to dry completely). 14. Resuspend the pellet in 50 L Tris-EDTA pH 8 or in 50 L water. 4.2 Amplification of pREMI-Z Isolated From Pexophagy Mutants 1. Genomic DNA (5-10 L) is digested in a total volume of 20 L with EcoRI, HindIII, XbaI, SacI, or BglII (enzymes that do not cut pREMI-Z) for 20-24 hours at 37 C. 2. Add 180 L ddH2O and 200 L Phenol/Chloroform/Isoamyl alcohol (25:24:1). 3. Vortex the solution for 30 seconds, then microfuge (14,000xg; 2 minutes). 51

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4. Remove the majority of the aqueous top phase and discard, avoiding the proteins at the aqueous:phenol interface. 5. To the aqueous phase add half a volume (200 L) of chloroform/isoamyl alcohol. 6. Again vortex the solution for 30 seconds then microfuge (14,000xg; 2 minutes). 7. Remove 180 L of the top phase (aqueous layer) and transfer to a clean tube. 8. Add 20 L of 3M sodium acetate (NaOAc) and 400 L 100% Ethanol. 9. Place the vial for 20 minutes in the -80 C freezer or in dry ice. 10. Microfuge (14,000xg; 2 minutes) and decant supernatant. 11. Wash the pellet with 0.5 mL cold 70% EtOH. 12. Microfuge (14,000xg; 2 minutes) and decant supernatant. 13. Air-dry the pellet (approximate ly 20 minutes; do not allow the pellet to dry completely). 14. Resuspend the pellet in 20 L water. 15. Incubate 5 L of digested genomic DNA in a total volume of 10 L with T4 DNA ligase. 16. Incubate 20-24 hours at 16 C. 17. Add 5 L of the ligation reaction to 100 L Rb-competent DH5 cells. 18. Place the solution on ice for 2 minutes, then 42 C for 90 seconds, then ice for 2 minutes. 19. Add 0.5 mL YETM and incubate for 1 hour at 37 C with gentle shaking. 20. Plate onto LB+Z. 21. Colonies are picked up and grown in liquid LB+Z 22. The pREMI-Z vector with flanking DNA is isolated using Qiagen miniprep spin columns. 52

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4.3 Sequencing Flanking Genomic DNA 1. The Big Dye sequencing reaction is done following routine procedures (Applied Biosystems) using M13 reverse and M13 (-20) primers. 2. Sequencing is done on an Applied Biosystems (ABI) 310 Prism Capillary Electrophoresis DNA Sequencer. 3. The disrupted gene is then identifi ed by BLAST analysis on the NCBI and S. cerevisiae databases. Notes 1. In some studies, the linearized pREMI-Z is de phosphorylated with calf intestinal phosphatase prior to electroporation (Mukaiyama et al., 2002). 2. The addition of a restriction enzyme during electroporation with the linearized pREMI-Z is believed to generate free chromosomal ends in the yeast genome for incorporation of the REMIZ plasmid. However, the number of transfor mations was only increased two-fold when investigated in H. polymorpha (van Dijk et al. 2001). Furthermore, doses of BamHI higher than 2 units had a detrimental affect on the efficiency of transformation. Inte restingly, only thirty percent of the integrations had occurred at a Ba mHI site and in many cases the BamHI site was not conserved because of nucleotide loss from the pREMI-Z vector (van Dijk et al. 2001). However, REMI mutagenesis reported for C. albicans and S. cerevisiae do require the addition of restriction enzymes (Sch iestl and Petes, 1991; Brown et al. 1996). 3. The integration of the pREMI-Z vector is st able for several generations of growth in nonselective YPD medium (van Dijk et al. 2001). A majority of the in tegrations occur at a single site within an open reading frame that can be easily characterized. However, there are some genomic integrations that coul d complicate the characterization of the mutant as well as the identification of the disrupted gene First, the pREMI-Z could inse rt into more than one gene. 53

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Although we have found that this is a rare ev ent, each disrupted gene would have to be characterized individually. Van Dijk, et al. have observed only two out of forty H. polymorpha transformants contained insertions into two genomic loci (van Dijk et al. 2001). Second, there could be multiple insertions of the pREMI-Z vect or into the same site. Although a single gene would be disrupted, the tandem insertion of pREMI-Z vectors would make it difficult to obtain any meaningful sequence data. Studies in H. polymorpha have shown that single copy integration occurred in 50% of the transformants, double copy inte gration occurred in 25% of the transformants, and 3 or more c opies of pREMI-Z occurred in 25% of the transformants (van Dijk et al. 2001). It appears that the pREMI-Z cassette formed multimers prior to integration since it occurred independent of the addi tion of restriction enzy mes. Finally, the insertion could cause a genomic deletion. These can be identified by sequencing the flanking DNA isolated with the pREMI-Z vector and noting that the sequence on both sides of th e pREMI-Z is not continuous. Finally, intergenic insertions are also difficult to interpret. 4. The REMI protocol has been used successfully in both P. pastoris and H. polymorpha to isolate genes (GSA, glucose-i nduced selective autophagy; PAZ, pexophagy zeocin-resistance; PDD, peroxisome degradation-deficient) essent ial for peroxisome degradation (see Table 1). The quantification of AOX activity has been th e screen of choice for pexophagy mutants in P. pastoris. A more detailed characterization of these mutants can be done by observing the vacuole morphology labeled with FM4-64 relative to the peroxisomes labeled with BFP-SKL or GFP-SKL (Kim et al., 2001; Stromhaug et al., 2001; Mukaiyama et al., 2002). However, a second screen has been used to identify pexophagy mutants in H. polymorpha (van Dijk et al., 2001). This qualitative screen requires the micr oscopic observation of th e prolonged presence of eGFP-SKL (eGFP protein which is C-terminally ex tended with a peroxisomal targeting signal of 54

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serine-lysine-leucine-COOH) in cells after the shift from YNM to YND medium. The REMI protocol has also been utilized to identify muta nts defective in peroxisome biogenesis (van Dijk et al. 2001). In general, these mu tants fail to thrive when grown on YNM plates. However, a more detailed characterization of the mutant rega rding protein sorting or aberrant number or size of peroxisomes can be done by microscopic visualization of eGFP-SKL (van Dijk et al. 2001). 5. Efficient lysing of the yeast is critical for accurate measurements of AOX activity. For the direct colony assay, cells were originally lysed by digestion of the cell wall with Zymolase 20T (0.25 mg/mL). However, this procedure yielde d variable results and we found that liquid nitrogen freezing and thaw ing was more efficient. For cells in liquid suspension, the cells are lysed by vortexing in the presence of glass beads. 6. It is possible that some genomic fragments c ontaining the pREMI-Z vector may be too large to be amplified in E. coli. However, we have been able to am plify vectors that are 10-12 kb. If the vectors are too large, then we s uggest using a different restricti on enzyme to digest the genomic DNA for recovery of the pREMI-Z. In principal, any enzyme that does not cut pREMI-Z would be appropriate. 7. The Zeocin antibiotic is sensitive to high salt concentrations. Normall y, the selection of yeast that had been electroporated cons ists of growth on minimal medi um plates (0.67% yeast nitrogen base without amino acids, 2% dextrose, 1M sorbito l, 0.4 mg/L biotin, 2% agar). However, we have found that Zeocin does not appear to be active in such medium. Therefore, we have utilized YPD plates with Zeocin. Furthe rmore, the selection of Zeocin-resistant E. coli is done in medium containing only 0.5% NaCl. 8. Restriction enzyme-mediated integration ha s been utilized successfully to generate mutants and identify genes essential for a num ber of cellular pathways in many different 55

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organisms including: Saccharomyces cerevisiae (Schiestl and Petes, 1991), Candida albicans (Brown et al. 1996), Coprinus cinereus (Granado et al., 1997), Lentinus edodes (Sato et al., 1998), Gibberella fujikuroi (Linnemannstons et al. 1999), and Aspergillus fumigatus (Brown et al. 1998) 56

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Table 2-1: Pexophagy genes identify by REMI. Pichia pastoris1 Hansenula polymorpha2 Characteristics References GSA10 PAZ1 PDD7 ATG13 Serine/threonine-kinase that complexes with Atg11 and Vac8 (Stromhaug et al., 2001; Mukaiyama et al., 2002; Komduur et al., 2003) PAZ2 HpAtg8 ATG8 Soluble protein that becomes conjugated to lipids (Mukaiyama et al., 2004; Monastyrska et al., 2005) PAZ3 ATG16 Component of Atg12-Atg5 complex (Mukaiyama et al., 2002) PAZ4 ATG26 UDP-glucose:sterol glucosyltransferase (Mukaiyama et al., 2002) PAZ5 GCN3 Translation initiation factor (Mukaiyama et al., 2002) GSA9 PAZ6 PDD18 ATG11 Coiled-coil domain found at vacuole surface (Kim et al., 2001; Klionsky et al., 2003) GSA11 PAZ7 ATG2 Peripheral membrane protein that interacts with Atg9 (Stromhaug et al., 2001; Mukaiyama et al., 2002) PAZ8 ATG4 Cysteine proteinase (Mukaiyama et al., 2002) GSA14 PAZ9 ATG9 Integral membrane protein associated with organelles juxtaposed to the vacuole (Stromhaug et al., 2001; Mukaiyama et al., 2002) PAZ10 GCN1 Regulates translation elongation (Mukaiyama et al., 2002) PAZ11 GCN2 Regulates translation initiation (Mukaiyama et al., 2002) PAZ12 ATG7 E1-like enzyme responsible for the conjugation of Atg12 to Atg5 and Atg8 to lipids (Mukaiyama et al., 2002) GSA19 PAZ13 PDD19 VPS15 Membrane-anchored Serine/threonine-kinase with WD40 domains (Stromhaug et al., 2001; Mukaiyama et al., 2002) GSA15 PAZ14 PEP4 Endopeptidase (Mukaiyama et al., 2002) PAZ16 ATG24 Sorting nexin with PX domain (Ano et al., 2005) PAZ19 GCN4 Transcriptional activator GSA12 ATG18 WD40 protein (Guan et al., 2001) GSA20 ATG3 E2-like enzyme responsible for the conjugation of Atg8 to lipids PDD13 MPP1 Zn(II)2Cys6 Transcription Factor (Leao-Helder et al., 2003) PDD15 ATG21 WD40 protein (Leao-Helder et al., 2004) PDD20 VAM3 Vacuolar t-SNARE 1 GSA, glucose-induced selective aut ophagy; PAZ, pexophagy zeocin-resistance. 2 PDD, peroxisome degradation-deficient. 3 A unified gene nomenclature of autophagy and autophagy-related processes such as pexophagy has been standardized across species to be ATG (autophagy-related) for the gene names (Klionsky et al. 2003). 57

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Figure 2-1: Insertional mutagenesis by restric tion enzyme-mediated integration of linearized pREMI-Z. GS115 cells are transformed by the insertion of linearized pREMI-Z (NCBI accession number AF282723) into the BamHI or DpnII site of a putative GSA gene. Zeocin-resistant cells that are defective in pexophagy are isolated and the GSA/PAZ ge ne identified by excising the pREMI-Z and flanking genomic D NA by restriction digest. 58

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Figure 2-2: Insertion of pREMI-Z into the ge nomic DNA of REMI mutants as visualized on Southern blots. Genomic DNA isolated from REMI mutants was digested with EcoRI, the resulting DNA fragments separated on agaros e gels, and those DNA fragments containing pREMI detected on Southern blots using biot inylated probes prepared from pREMI-Z. Figure 2-3: The direct colony a ssay of AOX activity in REMI-mut ated cells grown on YNM (A) or adapted from YNM to YND (B). The arrow indicates a colony defective in degrading AOX. 59

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Figure 2-4: Glucose-induced degr adation of AOX in REMI mutants. Parental GS115, autophagy-defective atg7 mutants, vacuole-defective pep4/prb1 mu tants, and REMI mutants (R2, R5, R8, R10, R12, R19, R22, and R115), were gr own in YNM medium and then switched to YND medium. At 0h and 6h, the cells were lyse d and AOX activity determined. Only 10% of the AOX remained at 6h of glucose adaptation in GS115, while 90% was present in atg7 and pep4/prb1 mutants. The pexophagy defect in the REMI mutants was not as severe as that observed in the atg7 mutants with 40-70% of the AOX activity remaining. 60

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CHAPTER 3 PPATG9 TRAFFICKING DURING PEXOPHAGY2 Introduction Cellular activities are regulated and mainta ined as a well-controlled balance between protein synthesis and degradation of essential proteins and enzyme s. Protein lifetimes can last from minutes to days with a majority of th e cellular proteins bei ng degraded in a lytic compartment such as the lysosome or vacuol e. Autophagy is the primary pathway for the delivery of endogenous proteins a nd organelles to this compartment (Levine and Klionsky, 2004; Klionsky, 2005). There exist a number of autophagic pathways in yeast: microand macroautophagy, microand macro-pexophagy, cytopl asm-to-vacuole targeting pathway (CVT)3, and vacuole import and degradation pathway. Th e CVT pathway is constitutive while the other pathways are regulated by envir onmental signals. For example, autophagy is enhanced when cells are deprived of nutrients such as amino acids and glucose while pexophagy is activated when methylotrophic yeasts ad apt from growth on methanol medi um to growth on glucose or ethanol medium. Autophagy sequesters cytosolic proteins and organell es nonselectively while pexophagy is responsible for the selective degradation of peroxisomes. Despite their differences, these pathways share a number of common molecular events. For example, Atg7/Gsa7/Apg7, Atg1/Gsa10/Apg1/Aut3, Atg2/Gs a11/Apg2, Atg18/Gsa12/Cvt18, and Vps15 are required for 2 Article reprinted with permission from publisher: Chang, T., L.A. Schroder, J.M. Thomson, A.S. Klocman, A.J. Tomasini, P.E. Strmhaug, and W.A. Dunn, Jr. (2005) PpATG9 encodes a novel membrane protein that traffics to vacuolar membranes which sequester peroxisomes during pexophagy in Pichia pastoris. Mol Biol Cell.16(10):494153. 3 The abbreviations used are: AOX, alcohol oxidase; FM 4-64, N-(triethlyammoniumpropyl)-4-(pdiethylaminophenylhexatrienyl) pyridinium dibromide; TCA, trichloroacetic acid; GFP, green fluorescent protein; BFP, blue fluorescent protein; mRFP, monomeric red fluorescent protein; GSA, glucose-induced selective autophagy; REMI, restriction enzyme-mediated integration; CVT, cytoplasm-to-vacuole targeting; Atg9-PC, Atg9 peripheral compartment; PVS, perivacuolar structure; MIPA, micropexophagy-specific membrane apparatus; PAS, pre-autophagosome structure. 61

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pexophagy, autophagy and CVT pathways, while At g11/Gsa9/Cvt9 and Vac8 are essential for pexophagy and CVT pathways, but not autophagy (Stasyk et al. 1999; Yuan et al., 1999; Scott et al. 2000; Guan et al., 2001; Kim et al., 2001; Stromhaug et al., 2001; Stromhaug and Klionsky, 2001; Wang et al., 2001; Huang and Klionsky, 2002; Abeliovich et al. 2003). Since many autophagy genes and their orthologues have been identified using different yeast models, the nomenclature for these genes had become co nfusing. Therefore, we will utilize the ATG4 nomenclature for those genes uniquely essential for autophagy (Klionsky et al. 2003). Pichia pastoris is able to synthesize enzymes to assimilate methanol for energy and growth. These enzymes such as alcohol oxidase (AOX) are housed primarily in the peroxisomes (Tuttle et al. 1993). When these cells adapt from meth anol to ethanol or glucose, the now superfluous peroxisomes are rapi dly degraded by autophagic even ts. During ethanol adaptation, peroxisomes are individually and selectively se questered into autophagosomes that then fuse with the vacuole. This process is call ed macropexophagy. A second venue for selective peroxisome degradation occurs during gluc ose adaptation. Micropexophagy proceeds by a mechanism whereby the vacuolar membrane inva ginates, resulting in protrusions that surround and engulf the entire cluster of peroxisomes (Tuttle et al. 1993; Sakai et al. 1998). Both pathways result in the degradation of peroxisomes within the vacuole by proteolytic enzymes. Pichia pastoris is a particularly good genetic mo del for the study of pexophagy because of its ability to rapidly and selectively degr ade peroxisomes (i.e., AOX) when adapting from methanol to glucose medium. The sequestration and degradation of peroxisomes during glucose adaptation is completed within 6 h (Tuttle and Dunn, 1995). Based on the movements of the vacuole that can be easily observed by fluor escence and electron micr oscopy, glucose-induced 4 We will conform to the ATG nomenclature; all the autophagy-related (ATG) genes will contain a genus and species prefix (e.g., PpAtg for the P. pastoris and ScAtg for the S. cerevisiae homologues). 62

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micropexophagy has been described as a multistage process that include s: signaling events (vacuole round), early sequestrati on events (vacuole indented), in termediate sequestration events (vacuole indented with short arm-like processes) late sequestration events (vacuole indented with long arm-like processes), and vacuole degr adation (vacuole with au tophagic bodies) (Sakai et al. 1998; Stromhaug et al., 2001; Mukaiyama et al., 2002). We and others have utilized genetic screens to identify a number of unique proteins required for pero xisome degradation in Pichia pastoris (Tuttle and Dunn, 1995; Sakai et al. 1998; Stromhaug et al., 2001). In order to better understand the functions of these proteins, we examined the vacuole morphology upon glucose-induced pexophagy. For example, PpVps15 (Paz13), PpAtg11 (Gsa9), and PpAtg18 (Gsa12) appear to act at early sequestration events (Stasyk et al. 1999; Guan et al., 2001; Mukaiyama et al., 2002), PpAtg2 (Gsa11) and PpAtg7 (Gsa7/Paz12) at intermediate sequestration events (Yuan et al., 1999; Stromhaug et al., 2001), and PpAtg1 (Paz1/Gsa10) at late sequestration events (Mukaiyama et al., 2002). Many of these proteins are structurally and functionally homologous to thos e proteins essential for aut ophagy and CVT pathways in S. cerevisiae (Habibzadegah-Tari and Dunn, 2003; Klionsky et al. 2003). Furthermore, structural homologues of these proteins can be found in various inverteb rates and vertebrates including humans (Klionsky et al., 2003). In this study, we have sequenced and char acterized a unique membra ne protein that is required for both pexophagy and autophagy in P. pastoris Based on vacuole morphology during glucose adaptation, we project th at PpAtg9 is required for an ear ly sequestration event. In growing cells, this protein local izes to unique foci near the cell periphery. During glucoseinduced pexophagy, PpAtg9 traffics from these vesicles to perivacuolar struct ures that appear to adjoin the vacuole and then to the sequestering membranes that arise from the vacuole. The 63

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trafficking of PpAtg9 requires PpAtg7, PpAtg11, PpAtg2, and PpVps15, but not PpAtg1, PpAtg18, or PpVac8. Furthermore, our data suggest that the transfer of PpAtg9 to the vacuole is a prerequisite for the formation of those sequestering membranes that engulf the peroxisome for incorporation into the vac uole for degradation. Experimental Procedures Yeast Strains And Media The yeast strains used in this study are liste d in Table 1 and were routinely cultured at 30C in YPD (1% Bacto yeast extract, 2% Bacto peptone, and 2% dextrose). P. pastoris was grown in YNM (0.67 % yeast nitrogen base, 0.4 mg/l biotin, and 0.5% methanol) to induce peroxisome biogenesis. The degradation of pe roxisomes was induced when cells grown in YNM were transferred to YND (0.67 % yeast nitrogen base, 0.4 mg/L biotin, and 2% glucose) or YNE (0.67 % yeast nitrogen base, 0.4 mg/L biotin, and 0.5% ethanol). Nitrog en starvation medium contained 0.17 % yeast nitrogen base (without amino acids and NH4SO4) and 2% glucose. All media contained 2% agar when made as plates. Histidine or argini ne or both were added at 40 g/ml when needed. Vector amplifica tion was done in E. coli (DH5 ) cultured at 37C in LB (0.5% Bacto yeast extract, 1% Bacto tryptone, and 1% Na Cl) with ampicillin (100 g/ml). Zeocin was added at 25 g/ml when culturing DH5 and 100 g/ml when culturing P. pastoris Yeast Transformation Cells grown overnight in YPD to an optical density (OD600) of 1.0 were harvested and treated with 10 mM DTT in YPD containing 25 mM HEPES, pH 8, for 15 min at 30C. The cells were washed twice in ice-cold water and once in 1 M sorbitol and then resuspended into 1 M sorbitol. Cells (40 l) were mixed with 0.2-1 g of linearized vector and transferred to a 0.2 cm gap cuvette (Bio-Rad, Hercules, CA), and the DNA was introduced by electroporation at 1.5 kV, 64

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25 F, 400 (Gene Pulser, Bio-Rad Corp.). The cells transformed with vectors containing the His4 gene were transferred to plates containing 0.67% yeast nitr ogen base without amino acids, 2% glucose, 1M sorbitol, 0.4 mg/L bio tin, and 2% agar a nd incubated at 30 C for 3 5 d before colonies appeared. Cells transformed with vector s containing the zeocin resistance gene (zeoR) were grown on YPD with 100 g/ml Zeocin. Isolation of GSA Mutants and Cloning of GS A Genes By Restriction Enzyme Mediated Integration (REMI) Mutagenesis Mutagenesis was performed by randomly insert ing the pREMI-Z vector (provided by Dr. Ben Glick, Univ. of Chicago) th at contained the Zeocin resist ance gene into the genome of P. pastoris as previously described (Stromhaug et al. 2001). Those gsa mutants caused by disruption of gene expression were identif ied by direct colony assays (Stromhaug et al., 2001). The site of insertion of the pREMI-Z and iden tification of the disrupted gene was done as described (Stromhaug et al., 2001). Based on the genomic sequences around the pREMI-Z insertion site that was isolated along with the pREMI-Z vector from the R19 mutant, GSA14 was cloned and sequenced from genomic DNA using a linker-mediated PCR method previously described (van Der Wel et al. 2001). We were able to completely assemble the GSA14 gene (NCBI accession number AY075105) and show it en codes a protein homologous to ScAtg9 of Saccharomyces cerevisiae, using the adapted ATG nomenclature for those genes uniquely essential for autophagy (Klionsky et al. 2003). A search of the National Center for Biotechnology Information (NCBI) database re vealed structural homologues of PpAtg9 and ScAtg9 in Schizosaccharomyces pombe (NP_596247), Arabidopsis thaliana (NP_180684), Neurospora crassa (XP_331198), Drosophila melanogaster (NP_611114), Caenorhabditis elegans (NP_503178), and Homo sapiens (NP_076990) (Yamada et al., 2005). The alignment of these proteins reveals a large cen tral region (190-679 residues) of homology containing at least 65

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five putative transmembrane domains. The pREM I-Z disruption of PpAtg9 occurred after the first putative transmembrane domain at aspartic ac id 245. Using this approach, we have isolated R2, R12, R13 and R22 mutants. The R2 ( his4 Ppatg18-1 ::zeoR) and R13 ( his4 Ppatg11-1:: zeoR) mutants have been described elsewhere (Guan et al. 2001; Kim et al., 2001; Stromhaug et al., 2001). The R12 mutants had the pREMI-Z inserted into the PpATG1 gene loci. The R22 ( his4 Ppatg2-2 :: zeoR) and WDK011 ( his4 Ppatg2:: zeoR) mutants have been characterized previously (Stromhaug et al., 2001). The Ppatg7/gsa7 mutants have been previously described (Yuan et al., 1999). The null mutant of PpVps15 was provided by Dr. J. Cregg (Keck Graduate Institute) (Stasyk et al. 1999). PpVAC8 was cloned from genomic DNA using degenerate primers combined with linker-mediated PCR (van Der Wel et al. 2001). The entire gene was sequenced and assembled (NCBI accession number AY886543). A Ppvac8 null mutant was constructed by replacing the entire gene ( bp through 1669 bp) with the zeoR gene driven by the TEF1 promoter. The null mutants were selected on YPD plates containing 100 g/ml Zeocin and replica-plated to YNM plat es. Null mutants were identified by direct colony assay (see below) and verified by P CR using primers flanking PpVAC8 and within the zeoR gene (data not shown). Measurements Of Alcohol Oxidase (AOX) And Endogenous Protein Degradation The direct colony and liquid medium assays to detect and measure the degradation of peroxisomal AOX was performed as previously described (Stromhaug et al., 2001). Briefly, cells were grown in YNM for 40 h. At which ti me, glucose (2%) or ethanol (0.5%) was added. Aliquots of cells (2 ml of OD600 = 1.4) at 0 and 6 hours of glucose or ethanol adaptation were pelleted and resuspended in 1 ml 20 mM Tris, pH 7.5, containing 50 mM NaCl, 1 mM EDTA, 1 mM PMSF, 1 g/ml pepstatin A, and 0.5 g/ml leupeptin. The cells were then lysed by 66

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vortexing in the presence of gla ss beads (425-600 microns). The gl ass beads and cellular debris were removed by centrifugation and AOX measured (Tuttle and Dunn, 1995; Yuan et al. 1999). The degradation of endogenous pr oteins during nitrogen starvati on was performed as described previously (Stromhaug et al., 2001). Briefly, cellular protei ns were radiolabeled with 14C-valine for 16 h and the cells switched to nitrogen st arvation medium containing 0.17 % yeast nitrogen base (without amino acids and NH4SO4) and 2% glucose and supplemen ted with 10 mM valine. The rates of protein degradation were calculated from the slopes of the linear plots of TCAsoluble radioactivity over 2-24 h of chase. Western Blot Analysis Cells (2 mls) from cultures gr own to an optical density (OD600) of 1.4 were collected by centrifugation and prepared for SDS-PAGE as previously described (Tuttle and Dunn, 1995). The cells were lysed in 67 mM Tris, pH 6.8, 2% SDS, 10% glycerol, 0.1% bromophenol blue, 1.5% DTT solution and 2 l of a proteinase inhibitor cock tail (200 mM phenylmethylsulfonyl fluoride, 14.5 mM pepstatin A, 10.5 mM leupeptin in DMSO) by vortexing with glass beads. The proteins were separated by SDS-PAGE and then transferred to nitrocellulose by Trans-Blot Semi-Dry Transfer Cell (Bio-Rad Laboratories, Hercules, CA) for 1 h. The blots were blocked in 5% nonfat dried milk in PBS and then incubated with mouse anti -AOX or rabbit anti-HA antibody (Covance Inc., Princeton, NJ). Following incubation with secondary goat anti-mouse antibody conjugated with HRP (Covance Inc., Prin ceton, NJ), the blots were washed and HRP detected using ECL-plus (Amersham, Piscat away, NJ) and quantified using the Typhoon 9400 laser scanner (Molecular Dynamics, Sunnyvale, CA). Construction of PpAtg Expression Vectors The gene for the green fluorescent protein (G FP) was inserted behind the glyceraldehyde 3-phosphate dehyrogenase (GAPDH) promoter into the EcoRI site of pIB2 (Sears et al. 1998). 67

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The resulting expression vector, pPS55, was then used to construct the GFP fusion protein of PpAtg9 being expressed by the constitutive and glucose-inducible GAPDH promoter. PpATG9 was amplified from genomic DNA by PCR with EnzyPlus polymerase (Enzypol, Ltd., Boulder, CO) using a forward primer of 5CTCTCACATTGTCGGTACCATGCATAAGAATAACACGAC -3 that contained a KpnI site. The reverse primer 5-GTTTTGGACTCGAGGGTA CTAATGCTTCATT-3 contained a XhoI site. PpATG9 was inserted behind the GFP gene in pPS55. A second expression vector called pTC2 was created by inserting PpATG9 with its endogenous promoter in front of GFP in pWD3 whereby GFP had been inserted into th e SphI site of pIB1 (Sears et al. 1998). PpATG9 was amplified from genomic DNA by PCR using a forward primer 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 that contained KpnI site and a reverse primer 5-CAAGATCTATGCTCGAGAACAAAT AATGCCTTATGCTGTTGACTAA-3 with a XhoI site. This product was then inserted into the KpnI and X hoI sites of pWD3, the resulting fusion construct verified by sequencing, a nd the vector used to transform R19 ( his4 atg9:: zeoR) cells. pTC3 was made using PpATG9 with a GAPDH promoter in place of the PpATG9 endogenous promoter by amplifying it from genomic DNA using a forward primer 5CTCTCACATTGTCGGTACCATGCATAAGAATAACACG AC-3 containing a KpnI site and a reverse primer 5-CAAGATCTATGCTCGAGAACAAATAATGCCTTAT GCTGTTGACTAA-3 with a XhoI site. It was inse rted in front of the GFP gene of pWD4, which had been inserted into the SphI site of pIB2 (Sears et al. 1998). Two additional expression vectors were made by inserting the PpATG9 gene behind the GFP and mRFP genes that were inserted into the EcoRI site of pGAPZ (Invitrogen, San Di ego). The gene for mRFP was kindly provided by Dr. R.Y. Tsein (University of Califor nia at San Diego) (Campbell et al. 68

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2002). PpATG9 was amplified from genomic D NA using a forward primer 5CTCTCACATTGTCGGTACCATGCATAAGAATAACACG AC-3 containing a KpnI site and a reverse primer 5-CATTATTATTCAAGACCGCG GAATGAAAA-3 with a SacII site. The PCR product was then cut with KpnI and SacII and inserted into pGAPz-GFP and pGAPz-RFP resulting in pWD17 and pAJM6 vectors, respectively. GFPPpatg9( N) lacking the N-terminus upstream of the first tran smembrane region and GFPPpatg9( L3) lacking a highly conserved third loop between the 3rd and 4th transmembrane regions wa s constructed by PCR. A Ppatg9 gene product lacking M1 through Y211 was am plified from genomic DNA by PCR using a forward primer 5-CGACTATGGTACCGGAAATG GATTCAA-3 with a KpnI site and a reverse primer 5-GTTTTGGA CTCGAGGGTACTAATGCTTCATT-3 with a XhoI site. The resulting gene was inserted into the K pnI and XhoI sites of pPS55. A second Ppatg9 gene product lacking T471 through A523 was construc ted by independently amplifying the gene fragments upstream and downstream of the deletion. The upstream region of the PpATG9 gene was amplified using a forward primer of 5CTCTCACATTGTCGGTACCATGCATAAGAATAACACG AC-3 that contained a KpnI site and a reverse primer of 5-CCGCAGGATTA AGCTTAAAATCGTAAAAA A-3 containing a unique HindIII site. The dowstream region of the PpATG9 gene was amplified using a forward primer of 5-CTACAACGAAGCTTCTGAAGTTCATCATGT-3 cont aining a HindIII site and a reverse primer 5-GTTTTGGACTCGAGGGTA CTAATGCTTCATT-3 containing a XhoI site. These two fragments were digested with the appropriate restriction enzymes and ligated into the KpnI and XhoI sites of pPS55. The expression vector, pWD21, contained PpAtg8 with a GFP at its N-terminus. PpAtg8 was amplified from genomic DNA by PCR using a forward primer of 5-CCATGAATCCATGCGATCGCAATTTAAAGACGAACA-3 containing an 69

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EcoRI site and a reve rse primer of 5GGCACTACTCGAGTTATTATTCAATCTCCTCAACACCT GGAA-3 containing a XhoI site. PpAtg8 was inserted behind the GFP gene in pPS55. The final expression vector, pASK1, contained PpAtg9 with an HA tag at its N-term inus driven by the endogenous promoter of PpATG9 This construct was made by PCR of the PpATG9 promoter and open reading frame separately, which was then followed by a PCR amplification of the entire gene including promoter. The promoter was amplified using a forward primer of 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 contai ning a KpnI site and a reverse primer of 5-GCGTAATCTGGAACATCGTATGGATACATTCAATCGAC AATGTGAGAGATTCAGTGAAG-3 containing the HA tag. The open reading frame was amplified using a forward primer of 5TGTATCCATACGATGTTCCAGATTACG CGATGCATAAGAATAA CACGACATTTTTAT CC-3 containing an HA tag (M YPYDVPDYA) insertion in front of the PpAtg9 start codon and a reverse primer of 5-GTCAGACTCCAAAC TCGAGTTCATTTTCAA-3 containing a XhoI site. Finally, the entire gene with promoter was amplified from the above products by PCR using forward 5-GCAGGCTAGGGTACCGGT ACTGGCACATT-3 and reverse 5GTCAGACTCCAAACTCGAGTTCATTTT CAA-3 primers. The resulting product was cut with KpnI and XhoI and inserted into pIB 1. The CoxIV signal sequence plus the upstream ADH1 promoter was excised from pCC4 by cutting with EcoRV and XbaI (Campbell and Thorsness, 1998). The fragment was then inserted into the SmaI and SpeI sites upstream of the GFP gene of pWD3, which had been constructed by inserting the GFP gene into the SphI site of pIB3 (Sears et al. 1998). The resulting expression vect or, pAJM3, produced a GFP protein that was targeted to the mitochondria. The constr uction of pPS55-G12, pPS64, and pPS69 vectors 70

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have been previously described (Guan et al., 2001; Kim et al., 2001; Stromhaug et al. 2001) and pPOP-S7-GFPX3, pPOP-SEC13-GFP, and pIB2 -DsRED-HDEL were provided by Dr. Ben Glick (Univ. of Chicago) (Bevis et al. 2002). For transformation, these vectors were first linearized by cutting either within the HIS4 gene with StuI or SalI or within the PpATG9 gene with SacI. Fluorescence Microscopy And FM 4-64 Labeling Cells expressing GFP or RFP fu sion proteins were grown in either YPD for 24 h or YNM for 20 h. Cells grown on YNM medium were then transferred to YND or YNE for 1 to 4 h. FM 4-64 (Molecular Probes, Eugene, OR) was added to a final concentration of 20 g/ml and the cells incubated for 2-12 h. The cells were washed of unbound FM 4-64 and examined immediately using a Zeiss Axiophot fluorescen ce microscope. Image capture was done using SPOT camera (Diagnostics Instruments, Inc., St erling Heights, MI) in terfaced with IP Lab software. Electron Microscopy The cellular ultrastructure of Ppatg9 mutants was examined as previously described (Tuttle and Dunn, 1995). Briefly, cells grown in YNM or grown in YNM and adapted to YND or YNE were harvested by centrifugation, washed in water, and fixed in 1.5% KMnO4 in veronal-acetate buffer (28 mM sodium acetate, 28 mM sodium barbital, pH 7.6) for 20 min at 22 C (Veenhuis et al. 1983). The specimens were dehydrated with increasing concentr ations of ethanol and infiltrated with POLY/BED 812 (Polysciences, Inc., Warrington, PA) with accelerator 2,4,6Tri(dimethylaminomethyl) pheno l (DMP-30, Polysciences, Inc.) for 2 d at 22 C under vacuum. After polymerization of the re sin, the samples were sectioned and examined using a JEOL 100CX transmission electron microscope. 71

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Results Ppatg9 Mutants Are Defective In Glucose-Induced Pexophagy And Starvation-Induced Autophagy Pichia pastoris can assimilate methanol for growth by synthesizing peroxisomal enzymes such as alcohol oxidase (AOX). When these cells are switched from a medium containing methanol to one containing glucose, the peroxi somes are selectively se questered by the vacuole for degradation by a process called micrope xophagy (Tuttle and Dunn, 1995). In order to identify the molecular components of this degr adative process, we developed a novel approach whereby genes can be randomly mutagenized in vivo by the integration of a vector containing a zeoR gene, pREMI-Z. Those Zeocin resistant ce lls that were unable to degrade AOX during glucose adaptation were identified by direct colony assay and verified by liquid medium assay (see Materials and Methods). Usi ng this approach, we have isolat ed a number of mutant strains based on their poor ability to degrade AOX during glucose adaptation. The gene mutated by the insertion of the pREMI-Z was sequenced and identif ied in each of our mutants (see Table 3-1). We then compared the inability of these mutant s to degrade peroxisome s to parental GS115 and to vacuole defective mutants that lack proteinase s A (pep4) and B (prb1) (Figure 3-1A). Within 6 h of glucose adaptation, over 90% of th e AOX was degraded by the GS115 cells. In comparison, only 10% of the AOX was degraded in mutants lacking proteinases A and B. Less than 60% of the AOX was degraded in R19 cells that contained a pREMI-Z insertion within the open reading frame of the PpATG9 gene. AOX degradation in these R19 cells was comparable to that observed for Ppatg1, Ppatg2, and Ppatg11 mutants, but higher th an the 10-30% degraded by Ppatg7, Ppatg18, Ppvps15 or Ppvac8 mutants. Next, we examined the ability of R19 cells to degrade AOX during ethanol ad aptation (Figure 3-1B). We observed on Western blots a substantial loss of AOX protein when GS115 cells adapt from methanol to ethanol medium. 72

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However, no loss of AOX was detected in R 19 cells over 24 h suggesting that macropexophagy was suppressed in the cells lacking PpAtg9. Our previous studies have shown that a number of those REMI mutants defective in pexophagy were also defective in nitrogen starvation-enhanced proteolysis (Yuan et al. 1999; Guan et al., 2001; Stromhaug et al., 2001). Therefore, we next quantified protein degradation in R19 starved for amino acids (Figure 3-1C). When GS115 parental cells are starved for nitrogen and amino acids, the cellular protei n was degraded at a rate of 0.36 percent per hour. The rate of protein degradation in starved ce lls was significantly lower in cel ls lacking proteinases A and B consistent with this degradation being mediated by the vacuole. In cells lacking PpAtg11 or Vac8, cellular protein degradation was 70-90% of control suggesting these proteins have a minimal role in starvation-induced autophagy. To the contrary, we determined that the degradation of cellular proteins was reduced by 60% in Ppatg9 cells starved for amino acids and nitrogen. This value was comparable to the rates of degradation observed in Ppatg1, Ppatg2, Ppatg7, Ppatg18 and Ppvps15 mutants. Ppatg9 Is Essential For A Sequestration Event In Pexophagy We have shown that during glucose-induced micropexophagy the vacuole indents while sequestering membranes encircle multiple peroxisomes (Tuttle and Dunn, 1995). The sequestering membranes are labeled with FM 464, a dye that selectively labels vacuole membranes, suggesting that they likely form from the vacuole. A micropexophagic membrane apparatus (MIPA) containing PpAtg8 forms near the tips of the sequestering membranes. The MIPA is thought to assist in the fusion of th e sequestering membranes, thereby incorporating the peroxisomes into the vacuole for degrad ation. During ethanol-induced macropexophagy, individual peroxisomes are selectively incor porated into pexophagosomes whose membranes contain PpAtg8. The pexophagosome then fuses with the vacuole and its contents degraded. In 73

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order to better assess the site of blockage of pexophagy in the R19 cells, we examined the formation of the sequestering membranes a nd the MIPA during micr opexophagy and of the pexophagosome during macropexophagy. GS115 (Figure 3-2, A,C) and R19 (Figure 2, E, G, I, K) cells were grown in methanol-enriched YNM medium and then adapted to glucose-enriched YND (Figure 3-2, E, I) or ethanol-enriched YNE (Figure 3-2, G, K) medium. Cells were harvested at 3 h, fixed in pot assium permanganate, and prep ared for electron microscopy. During glucose adaptation, profiles of peroxiso mes engulfed by the vacuole were routinely observed in GS115 cells (Figure 2A). However, when R19 cells were adapted to glucose, the sequestering membranes appear to be absent, bu t instead the vacuole appeared only slightly indented (Figure 3-2E) with an occasional shor t arm-like projection (Fig ure 3-2I). Similar vacuole morphology has been reported for Ppatg11, Ppatg18 and Ppvps15 mutants (Stasyk et al. 1999; Guan et al., 2001; Kim et al., 2001)]. Next, we examined whether the MIPA was forming in R19 cells. WDY70 cells expressing BFP-SKL and GFP-PpAtg8 were adapted from YNM to YND for 2 h. At this time, PpAtg8 was lo calized to the MIPA and foci adjacent to the peroxisomes (Figure 3-2B). In the absen ce of PpAtg9, the MIPA was absent and PpAtg8 localized solely to foci (Fi gure 3-2, F, J). During ethanol adaptation, peroxisomes within pexophagosomes were observed by electron (Fig ure 3-2C) and fluorescence (Figure 3-2D) microscopy. In WDY70 cells, the PpAtg8 was localized almost exclusively to pexophagosomes (Figure 3-2D). Pexophagosomes were absent in cells lacking PpAtg9 (Figure 3-2, G, H, K, L), and PpAtg8 was localized to multiple foci that ap pear within and about the peroxisome cluster (Figure 3-2, H, L). The data dem onstrate that PpAtg9 is essential for the formation of sequestering membranes that arise from the vacuole and the assembly of the MIPA during micropexophagy and of the pexophagosome during macropexophagy. 74

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Cellular Localization Of Ppatg9 In Growing Cells Next, we examined the cellular distribution of PpAtg9 in cells grown in glucose medium. This was done by constructing the following ex pression vectors: GFP-PpAtg9 behind the GAPDH promoter in pIB2 (pTC1), PpAtg9GFP behind the endogenous PpAtg9 promoter (pTC2), GFP-PpAtg9 behind the GAPDH promot er in pGAPz (pWD17), and mRFP-PpAtg9 behind the GAPDH promoter in pGAPz (pAJM6). PpAtg9-GFP (data not shown), GFP-PpAtg9 (see Figure 3-8) and mRFP-PpAt g9 (see Figure 3-8) were functional as determined by their ability to rescue R19 cells. When these cells were grown in YPD, GFP-PpAtg9 (Figure 3-3A) localized to one or more foci or structures distributed about the cell peripher y. In order to better define the nature of these structures, we co-expressed GFP-PpAtg9 with DsRed-HDEL and mRFP-PpAtg9 with CoxIV-GFP in GS115 cells. Some structures containing GFP-PpAtg9 appeared to be in close association with endoplasmic reticulum identified by DsRed-HDEL (Figure 3-3C, arrow) and mitoc hondria identified by CoxIV-GFP (F igure 3-3D arrow). We next examined whether these structures contained Sec 13 (intermediate compartment) or Sec7 (Golgi apparatus) by co-expressing mRFP-PpAtg9 with Sec13-GFP and Sec7-GFPx3. The results demonstrate that PpAtg9 does not co-localize with either Sec13 (Figure 3-3E) or Sec7 (Figure 33F). In S. cerevisiae the peripheral vesicles containing ScAtg9 do not colocalize with any defined organelle markers for the Golgi appara tus, endosomes, or the endoplasmic reticulum (Noda et al., 2000; Kim et al. 2002; Reggiori et al. 2004). Therefore, we will refer to these unique peripheral structures collectively as th e Atg9 peripheral compartment (Atg9-PC). Cellular Trafficking Of Ppatg9 During Glucose-Induced Pexophagy The trafficking of PpAtg9 during glucose-i nduced pexophagy was visual ized in TC3 cells expressing BFP-SKL behind the AOX promoter and GFP-PpAtg9 behind the GAPDH promoter (Figure 3-4). When TC3 cells were grown in YNM, GFP-PpAtg9 localized to one or more 75

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structures, described above as the Atg9-PC (Fi gure 3-4A). Upon adapting these cells from YNM to YND for 3 h, GFP-PpAtg9 localized to multiple st ructures of differing si zes and shapes that were positioned at the vacuole surface labeled with FM 4-64 and to the sequestering membranes that could be seen flanking the peroxisomes labeled with BFP-SKL (Figure 3-4). We refer these structures that appear as dots and patches as perivacuolar structures (PVS). The PVS were routinely observed near those sites where th e sequestering membranes joined the vacuole suggesting they may function in the formation of these membranes from the vacuole. Indeed, the sequestering membranes containe d both PpAtg11 and PpAtg18 presen t at the vacuole membrane and stain with FM 4-64 suggesting that th ey originated from the vacuole (Guan et al., 2001; Kim et al. 2001). These results were not due to overe xpression of GFP-PpAtg9 that may be caused by the GAPDH promoter, since PpAtg9-GFP whose expression is regulated by the endogenous PpAtg9 promoter in TC14 cells showed a sim ilar distribution but w eaker signal (data not shown). In a given population of cells, the onset of micropexophagy is variable resulting in cells at differing stages of micropexophagy. Neverthele ss, we have presented a time course based on images obtained from 0-2h of glucose adaptation that best illustrates our understanding of PpAtg9 trafficking (Figure 3-4C). We show that during glucoseinduced pexophagy, PpAtg9 traffics from the Atg9-PC (arrowheads) to the PVS (arrows) and then to the vacuole membrane and those sequestering membranes (double arro wheads) that engulf the peroxisomes for degradation within the vacuole. We next examined the expression of PpAtg9 in cells during glucose-induced micropexophagy. This was done by constructing a PpAt g9 protein tagged at the N-terminus with an HA epitope. This construct along with the PpAtg9 promoter of about 400 bp upstream of the start codon was inserted into the pIB1 vector (Sears et al. 1998). The resulting vector was used 76

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to transform GS115 cells and the expression eval uated on Western blots using antibodies that recognized the HA tag. A single protein around 100 kDa was detected in cells expressing HAPpAtg9. This protein was absent in extracts of GS115 cells that were no t transformed (data not shown). At 4-6 h of glucose-induced pexophagy, the cellular levels of HA-PpAtg9 increased over two-fold relative to zero hour (Figure34D). Over the next 2 h, the cellular levels of HAPpAtg9 diminished. Similar results were obs erved when HA-PpAtg9 was expressed in R19 cells (data not shown). Cellular Trafficking Of Ppatg9 During Glu cose-Induced Pexophagy Requires Other Ppatg Proteins Many Atg proteins interact w ith each other, thereby influe ncing function and trafficking (Wang and Klionsky, 2003; Reggiori et al. 2004). In fact, the traffi cking of ScAtg9 appears to be regulated by a number of ScAtg proteins including ScAtg1 and ScAtg18 (Reggiori et al. 2004). Therefore, we investigated whether other PpAtg proteins we re required for the trafficking of PpAtg9 from the Atg9-PC to the PVS and sequestering membranes. We examined the trafficking of GFPPpAtg9 dur ing glucose-induced pexophagy in mutants defective in early ( Ppatg11, Ppatg18 and Ppvps15), intermediate ( Ppatg2 and Ppatg7), and late ( Ppatg1 and Ppvac8) sequestration events. These mutants were transformed by electroporation with pTC1 and GFP-PpAtg9 expression verifi ed by Western blotting and fl uorescence microscopy. When the resulting transformants were grown in YNM, GFP-PpAtg9 was found almost exclusively at the Atg9-PC (data not shown). After growth in YNM, the mutants expressing GFP-PpAtg9 were then adapted to glucose for 3 h. In Ppatg11 Ppatg18 and Ppvps15 mutants, the vacuoles were predominantly round with a slight indentation and no arm-like extensions were evident (Figure 5). GFP-PpAtg9 expressed in Ppatg11 and Ppvps15 cells was found predominantly at peripheral foci (Figure 5). In addition, the v acuole membrane was void of GFP-PpAtg9 in these 77

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mutants. In Ppatg18 cells, GFP-PpAtg9 localized to the PVS and to the vacuole membrane (Figure 3-5). The vacuoles in Ppatg7 and Ppatg2 mutants were indented with short extensions (Figure 3-6). In these cells, GFP-PpAtg9 was f ound in peripheral structures (arrowheads) and the PVS (arrows) located either at the vacuole or at the site where the sequestering membranes join the vacuole (Figure 3-6). Although the lo cation of the PVS appeared unaltered, the PVS morphology appeared to be smaller and less diffu se in these mutants than that observed in control TC3 cells (see Figure 3-4). Furthe rmore, GFP-PpAtg9 was not detected at the sequestering or vacuole membranes. During micropexophagy, the vacuoles in Ppatg1 and Ppvac8 mutants contained an extensive segmente d array of sequestering membranes that extended from the vacuole to almost completely surround the peroxisome cluster (Figure 3-7). GFP-PpAtg9 was found at the PVS and seque stering membranes (Figure 3-7). Domains Of Ppatg9 Required For Function And Trafficking We have demonstrated that PpAtg9 traffics from the Atg9-PC to the PVS where it appears to function in the forma tion of the sequestering membranes. In order to define those domains that are essential for PpAtg9 functi on, we have characterized the function and trafficking of PpAtg9 mutants lack ing specific peptide re gions. We first deleted the N-terminus (M1-Y221), a region of low homo logy when compared to its S. cerevisiae counterpart. Unlike wild-type GFP-PpAtg9, GFP-Ppatg9 N only partially rescued R19, suggesting this protein has limited function for glucose-induced micropexophagy (Figure 3-8A). However, during glucose adaptation, GFP-Ppatg9 N could be observed at the Atg9-PC (Figure 3-8B, arrowheads), PVS (Figure 3-8B, arrows) and vacuolar membranes s uggesting that the N-terminus is not essential for PpAtg9 trafficking. Next, we deleted a highly conserved loop (T471 A523) between two putative transmembrane domains. As seen for Ppatg9 N, Ppatg9 L3 was also unable to 78

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efficiently rescue the R19 phenotype (F igure 3-8A). However, unlike Ppatg9 N, Ppatg9 L3 was not present at the Atg9-PC, PVS or vacuole membrane but to a compartment morphologically similar to the endoplasmic reticul um (Figure 3-8C, large arrows). The data suggest that loop T471 A523 is essential for trafficking of PpAtg9 to the Atg9-PC. Perivacuolar Structures Contain Ppatg9 And Ppatg11, But Not Ppatg2 We have previously shown that Atg2 beco mes associated with peripheral structures during pexophagy and that this asso ciation requires PpAtg9 (Stromhaug et al., 2001). In addition, we have reported that PpAtg11 localizes to the vacuole membrane and to one or more structures associated with e ither the vacuole or the arm-lik e extensions that surround the peroxisomes (Kim et al., 2001). In this study, we have show n that the trafficking of PpAtg9 requires PpAtg2 and PpAtg11. Therefore, we co mpared the localization of mRFP-PpAtg9 with PpAtg2-GFP and PpAtg11-GFP in ce lls adapting from YNM to YND (Figure 3-9). We first compared the localization of mRFP-PpAtg9 with PpAtg18-GFP, which we have previously shown to be present at the vacuole and sequestering memb ranes. In cells undergoing micropexophagy, mRFP-PpAtg9 was present in the At g9-PC and at the PVC localized at sites where the sequestering membranes appear to exte nd from the vacuole (Figure 9, arrows). Unlike GFP-PpAtg9, mRFP-PpAtg9 was not present at the vacuole membrane, but instead was present within the vacuole that wa s delineated by PpAtg18-GFP and PpAtg11-GFP. One possible explanation is that the mRFP at the N-terminus of PpAtg9 is exposed to vacuolar enzymes and proteolytically removed from PpAtg9. However, the accumulation of mRFP within the vacuole persisted in cells lacking Pe p4 or Prb1 (Tomasini and Dunn, unpublished observations). The data reveal that PpAtg2 does not colocalize with the Atg9-PC nor the PVS. However, some of the PpAtg2 vesicles can be found in close proxim ity to the PVS (see Fig. 3-9 arrows). PpAtg11 79

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localized to the vacuole membrane and the PVS containing PpAtg9 (Figure 3-9, arrows), but did not colocalize with the peripheral structures of PpAtg9 (F igure 3-9, arrowhead). The data suggest that the Atg9-PC does not contain PpAtg2 and that th e peripheral compartment of PpAtg2 may be unique. Furthermore, we have shown that the PVS contains PpAtg9 and PpAtg11 but not PpAtg2 or PpAtg18. Ppatg9 Does Not Localize To MIPA or The Pexophagosome Finally, we examined whether mRFP-PpAt g9 colocalized with GFP-PpAtg8 at the MIPA, at the pexophagosomes, or at the perivacu olar foci referred to as pre-autophagosome structures (PAS) in S. cerevisiae. During glucose-induced mi cropexophagy, PpAtg8 was present at perivacuolar foci and at the MIPA (arro wheads) which did not c ontain PpAtg9 (Figure 310A). During ethanol-induced macropexophagy, PpAt g8 was found at perivacuolar foci and at the pexophagosome (arrowheads), while PpAtg9 locali zed to the Atg9-PC (small arrow) and the PVS (large arrow) (Figure 3-10B). PpAtg9 did not colocalize with PpAtg8 and was absent from the pexophagosome. To the contrary, PpAtg8 wa s not present at the Atg9-PC (Figure 3-10B, small arrow) or at the PVS (Fi gure 3-10, large arrows). The data suggest that PpAtg9 is not a component of the MIPA or the pexopha gosome and that the PVS lacks PpAtg8. Discussion Autophagy is an avenue for the degradation of proteins and organell es. This process is critical for cell survival during times of nutrient deprivation. Twenty-seven autophagy genes (ATG) have been identified using yeast models of autophagy (Klionsky et al. 2003). One such model is Pichia pastoris in which we can follow the molecular events required for the selective sequestration and vacuolar degradation of pe roxisomes by a process called pexophagy. When cells adapt from growth in methanol to glucos e, the cells respond by ac tivating the degradation of peroxisomes by pexophagy. During glucoseinduced pexophagy, sequestering arms extend 80

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from the vacuole to engulf multiple peroxiso mes. Upon homotypic membrane fusion of the arms, the peroxisomes are incorporated into auto phagic bodies within the vacuole where they are degraded. We have previously shown that PpAtg2 (Gsa11), PpAtg7 (Gsa7), PpAtg11 (Gsa9), and PpAtg18 (Gsa12) are required for the se questration events of micropexophagy (Yuan et al., 1999; Guan et al., 2001; Kim et al., 2001; Stromhaug et al., 2001). In this study, we show that PpAtg9 is required for an early sequestrati on event in micropexophagy, possibly the initial formation of the sequestration membranes that extend from the vacuole. In addition, the assembly of MIPA during mi cropexophagy and the formation of the pexophagosome during macropexophagy does not occur in cells lacking PpAtg9. PpAtg9 is an integral membrane protein with five possible transmembrane domai ns that traffics to the vacuole membrane becoming a component of the sequestration membrane s that appear to arise from the vacuole. PpAtg9 is not essential for cell viability, but appears to be st ructurally conserved throughout a number of plant, fungi, insect, and mammalian sp ecies. PpAtg9 is also necessary for starvationinduced nonselective autophagy in P. pastoris S. cerevisiae, and Arabidopsis thaliana and for the trafficking of Ape1 to the vacuole in S. cerevisiae (Noda et al., 2000; Hanaoka et al. 2002). In S. cerevisiae ScAtg9 resides in two populations of vesicles, one at the cell periphery and a second found adjacent to the vacuole. Thes e vesicles distribute as a single peak on sucrose gradients which can be resolved from the v acuole (Pho8), plasma membrane (Pma1), Golgi apparatus (Kex2), endoplasmic reticulum (Sec12), and endosomes (Pep12) (Noda et al. 2000). In addition, our results s uggest that those vesicles adjacent to the vacuole are not the prevacuolar compartment (Noda et al. 2000). The pre-autophagosome stru cture (PAS) appears as a single structure located at the vacuolar surface and is co mposed of a number of proteins such as ScAtg2 (Apg2), ScAtg8 (Aut7), ScAtg9 (Apg9), ScAt g11 (Cvt9), ScAtg17 (Apg17), ScAtg18 (Cvt18), 81

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ScAtg20 (Cvt20), and ScAtg24 (Cvt13) (Suzuki et al. 2001; Huang and Klionsky, 2002; Kim et al. 2002; Nice et al. 2002; Reggiori et al., 2004). The PAS is thought to be responsible for organizing the formation of the autophagosome and transferring ScAtg8 to the autophagosome membrane. However, the appearance of these structures does not change when cells are starved for amino acids, and other components of the PA S such as ScAtg9 do not appear to associate with autophagosomes (Noda et al., 2000; Kim et al., 2002). Those ScAtg9 vesicles not adjacent to the vacuole but more at the cell periphery differ from the PAS in that they do not contain ScAtg8 (Kim et al., 2002; Tucker et al. 2003). In S. cerevisiae, ScAtg9 appears to recycle between the peripheral vesicles and the PAS. Furthermore, the movements of ScAtg9 from the PAS to the peripheral vesicles require ScAtg1 independent of its kinase activity, ScAtg2, ScAtg18 and the PtdIns 3-kina se complex I which includes ScVps34, ScVps15, and ScAtg14 (Reggiori et al. 2004). In Figure 3-11, we have presented a worki ng model for the trafficking of PpAtg9 during glucose-induced pexophagy. In growing cel ls, PpAtg9 resides in the Atg9 peripheral compartment (Atg9-PC) that cons ists of one or more structures distributed near the cell periphery. These structures are neither intermedia te vesicles nor the Golgi apparatus since they lack PpSec13 and PpSec7, respectively. Howeve r, they occasionally c ontain the endoplasmic reticulum marker, mRFP-HDEL, while others are in close association with the endoplasmic reticulum suggesting these structur es may be a subcompartment of the endoplasmic reticulum. Furthermore, Ppatg9 L3 is not present at the Atg9-PC, but appears to be associated with the endoplasmic reticulum. However, additional studie s are needed to substa ntiate the relationship between the endoplasmic reticulum and the Atg9-PC ScAtg9 appears to recycle between the peripheral compartment and the PAS (Reggiori et al. 2004). Such recycling was not detected in 82

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P. pastoris although we have not analyzed this under the same conditions defined in S. cerevisiae. We have also shown that the PpAtg9-PC is sometimes associated with mitochondria. However, the functional significan ce of this location remains unc lear. Nevertheless, based on our findings and those from S. cerevisiae we suggest that the peri pheral vesicles containing PpAtg9 are a unique compartment. Upon the onset of glucose-induced pexophagy, PpAtg9 is recruited from the Atg9-PC to tw o or more perivacuolar struct ures (PVS) and then to those membranes that sequester the peroxisomes. The PVS is in many ways similar to the PAS defined in S. cerevisiae. For example, both contain PpAtg9 and PpAtg11 (see Fig. 3-10). Ano et al. have shown that similar periva cuolar spots contain PpAtg24 (Ano et al. 2005). Both PVS and PAS are located at the vacuol e and appear to be essential for organizing the formation of sequestering membranes. However, the PVS doe s not contain PpAtg2 or PpAtg8 and appears to be structurally different from the PAS. Ther e exist multiple foci of PVS of differing sizes situated about the vacuole and the sequestering membranes while the PAS usually consists of a single, well-defined structur e at the vacuole. PpAtg17, a component of the PAS in S. cerevisiae, resides in a single structure ju xtaposed to the vacuole (Ano et al. 2005). PpAtg24 colocalizes with PpAtg17 but is also found at multip le other perivacuol ar structures (Ano et al., 2005). PAS has been implicated in the formation of th e autophagosome from membranes of unknown origin, but its function remains unclear (Suzuki et al. 2001). We have demonstrated that the PVS is situated at sites where the sequestering membrane s appear to form from the vacuole and that PpAtg9 is transported from the PVS to thos e sequestering membranes that engulf the peroxisomes. Therefore, we proj ect that PVS may act as the site of assembly in the formation of the sequestering membranes that appear to originate from the vacuole because they stain with FM 4-64. It is unclear how these membranes are formed. Mukaiyama et al., suggest these 83

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membranes form by a septation of the vacuole (Mukaiyama et al., 2002). Our data suggest that the formation of these membranes requires protein synthesis and both PpAtg9 and PpAtg18. We have previously shown that these membrane s do not form in the presence of cycloheximide (Tuttle and Dunn, 1995). In this study, we show th at the cellular levels of PpAtg9 are increased during pexophagy suggesting a role for protein synthe sis. In addition, we have demonstrated that when PpAtg9 is absent or does not traffic to the PVS, the vacuole arms are either not present or short. Conversely, when PpAtg9 localizes to the PVS and the vacuole membrane but PpAtg18 is missing, the vacuole arms are absent. Furthermor e, we have shown that PpAtg9 lacking its Nterminus is sorted to the PVS, but the peroxi somes are not sequestered or degraded. This suggests that there exists a region within the Nterminus of PpAtg9 that is essential for its function once it arrives at the PVS. This domain and its function have yet to be defined and are currently under inve stigation. We have shown that PpAtg11 and PpVps15, but not PpAtg18, PpAtg2, PpAtg7, PpAtg1, or PpVac8, are essential to recruit PpAtg9 to the PVS. In S. cerevisiae, the overexpression of ScAtg11 appears to enhance the recruitment of ScAtg9 to the PAS (Kim et al. 2002). In cells lacking ScAtg1, ScAtg2, ScAtg14, or ScAtg18, ScAtg9 is found predominantly at the PAS either because trafficking to the PAS is enhanced or re cycling to the peripheral vesicles is suppressed (Reggiori et al. 2004). ScAtg11 and ScAtg18 are present at the PAS and vacuole surface, but the localization of ScVps15 has not yet been determined (Guan et al., 2001; Kim et al., 2001). From the PVS, PpAtg9 is transferred to the membranes of the vacuole and those sequestering membranes that extend from the vacuole. The apparent movement of PpAtg9 from the PVS to the sequestering membranes requires PpAtg2 a nd PpAtg7, but not PpAtg18, PpAtg1 or PpVac8. We have previously shown that during pexophagy PpAtg2 becomes associated with foci of 84

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unknown origin (Stromhaug et al., 2001). In this study, we show that these structures are situated juxtaposed to the PVS. The homo typic fusion of the sequestering membranes and formation of the autophagic body within the vacuole completes the sequestration of the peroxisomes. A membrane sac called MIPA (micropexophagy-specific membrane apparatus), which contains PpAtg8 and PpAtg26, forms between the tips of the sequestering membranes to presumably direct membrane fusion (Oku et al., 2003; Mukaiyama et al., 2004). In addition to PpAtg8 and PpAtg26, the assembly of MIPA re quires the lipidation of PpAtg8, which is mediated by PpAtg3, PpAtg4, and PpAtg7. We show here that the assembly of MIPA also requires PpAtg9. However, it is unclear whether PpAtg9 has a direct role in MIPA formation or if the assembly of MIPA requires the presence of the sequestering membranes. Our data suggest that late sequestration or fusi on events require PpAtg1, a serine-threonine protein kinase, and PpVac8, an Armadillo-repeat protein structurally homologous to ScVac8 that when anchored to the vacuole membrane by palmitoylation wi ll promote homotypic vacuole fusion (Wang et al., 1998). PpAtg24 is also essentia l for homotypic fusion of th e sequestering membranes (Ano et al. 2005). PpAtg24 is not requir ed for MIPA formation (Ano et al., 2005), but it is unclear whether PpAtg1 and PpVac8 are essential for the formation of the MIPA. In summary, we have characterized the f unctional role of PpAtg9 in glucose-induced micropexophagy. We have shown that PpAtg9 is esse ntial for the formation of the sequestrating membranes that engulf the peroxisomes for degradation within the vacuole. During micropexophagy, PpAtg9 traffics from a unique compartment of the Atg9-PC to the PVS juxtaposed to the vacuole where it then becomes associated with the vacuole and those sequestering membranes that engulf the peroxiso mes for degradation (see Fig. 3-11). We have also demonstrated that the trafficking of PpAtg9 requires PpAtg11, PpVps15, PpAtg2, and 85

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PpAtg7, but not PpAtg1, PpAtg18 or PpVac 8. We propose that upon the onset of micropexophagy, PpAtg11 recruits PpAtg9 to the PVS, which act as sites of formation of the sequestering membranes presumably by causing segm entation of the vacuole. These membranes then engulf the peroxisomes and eventually fuse with the assistance of PpAtg1 and PpVac8 to incorporate the peroxisomes into the vacuole for degradation. 86

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Table 3-1: Pichia pastoris strains Name Genotype Reference GS115 his4 (Cregg et al., 1985) PPF1 arg4 his4 (Yuan et al., 1997) SMD1163 his4 pep4 pr b1 (Tuttle and Dunn, 1995) DMM1 GS115::pDM1 (PAOX1BFP-SKL, zeoR) (Kim et al., 2001) WDY7 his4 Ppatg7 (Yuan et al., 1999) WDKO7 PPF1 Ppatg7 ::ARG4 (Yuan et al., 1999) WDKO11 GS115 Ppatg2:: zeoR (Stromhaug et al., 2001) R2 GS115 Ppatg18-1:: zeoR (Stromhaug et al., 2001) R12 GS115 Ppatg1-1:: zeoR (Stromhaug et al., 2001) R13 GS115 Ppatg11-1:: zeoR (Kim et al., 2001) R19 GS115 Ppatg9-1:: zeoR (Stromhaug et al., 2001) R22 GS115 atg2-1:: zeoR (Stromhaug et al., 2001) Ppvps15 PPF1 his4 Ppvps15 ::ARG4 (Stasyk et al., 1999) S7GFPx3 PPF12 arg4 his4::pPOP-S7-GFPX3 (PSec7 PpSec7-GFPx3) (Bevis et al., 2002) Sec13GFP PPF12 arg4 his4::pPOP-SEC13-GFP (PSec13 PpSec13-GFP) (Bevis et al., 2002) WDY53 arg4 his4 vac8 :: zeoR This study WDY70 DMM1 his4::pWD21 (PGAPDH GFP-PpAtg8, HIS4) This study WDY71 R19 his4::pWD21 (PGAPDH GFP-PpAtg8, HIS4) This study WDY75 AJM26 his4::pWD21 (PGAPDH GFP-PpAtg8, HIS4) This study WDY78 R19 his4::pAJM9 (PGAPDH mRFP-PpAtg9, HIS4) This study TC1 GS115 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC3 DMM1 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC4 WDY7 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC5 WDKO7 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC6 R13 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC7 R12 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC8 R22 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC9 R2 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC10 R19 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC11 PpVsp15 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC12 WDY53 arg4 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study TC14 R19 his4::pTC2 (PGSA14 PpATG9-GFP, HIS4) This study TC16 R19 his4::pTC3 (PGAPDH PpATG9-GFP, HIS4) This study TC19 WDKO11 his4::pTC1 (PGAPDH GFP-PpATG9, HIS4) This study ASK2 GS115 his4::pHA-G14 (PGSA14 HA-PpATG9, HIS4) This study ASK3 R19 his4::pHA-G14 (PGSA14 HA-PpATG9, HIS4) This study ASK4 R19 his4::pGFP-G14( L3) (PGAPDH GFP-PpAtg9 L3, HIS4) This study ASK5 R19 his4::pGFP-G14( N) (PGAPDH GFP-PpAtg9N, HIS4) This study AJM18 S7-GFPx3 arg4 his4::pAJM6 (PGAPDH mRFP-PpATG9, zeoR) This study AJM19 Sec13-GFP arg4 his4::pAJM6 (PGAPDH mRFP-PpATG9, zeoR) This study AJM22 GS115 his4::pPS69(PGAPDH GFP/HA-PpATG2, HIS4) This study AJM24 GS115 his4::pWD17 (PGAPDH GFP-PpATG9, zeoR) This study AJM25 AJM22::pAJM6 (PGAPDH mRFP-PpATG9, zeoR) This study 87

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Table 3-1 continued Name Genotype Reference AJM26 GS115 his4 ::pAJM6 (PGAPDH mRFP-PpATG9, zeoR) This study AJM27 AJM24 his4 ::pIB2-dsRED-HDEL (PGAPDH dsRFP-HDEL, HIS4) This study AJM32 AJM26 his4 :: pPS64 (PGAPDH GFP/HA-PpATG11, HIS4) This study AJM33 AJM26 his4 :: pPS55-G12 (PGAPDH GFP/HA-PpAtg18, HIS4) This study AJM44 AJM26 his4 ::pAJM3(PADH CoxIV-GFP, HIS4) This study 88

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Figure 3-1. Pexophagy and autophagy are defective in Ppatg9 mutants. Panel A: Wild type GS115, R12 ( Ppatg1), R22 ( Ppatg2-2), WDY7 ( Ppatg7), R19 ( Ppatg9), R13 ( Ppatg11-1 ), R9 ( Ppatg18), Ppvps15 ( Ppvps15), WDY53 ( Ppvac8), and SMD1163 ( pep4, prb1) cells were grown in YNM for 36 h. At that time, cells were switched to a medium containing 2% glucose. Aliquots were removed at 0 and 6 h of adapta tion, the cells lysed, and AOX activities measured as describe in Experimental Procedures. The data is expressed as a percentage of AOX remaining at 6 h relative to 0 h and represents the mean SE of 3 6 trials. Panel B: Wild type GS115 and R19 ( Ppatg9) cells were grown in YNM for 36 h and then switched to YNE medium. Aliquots were removed, the cells lysed, and AOX protein visualized by Western blotting as describe in Experimental Procedures Panel C: Wild type GS115, R12 ( Ppatg1), R22 ( Ppatg22), WDY7 ( Ppatg7), R19 ( Ppatg9), R13 ( Ppatg11-1 ), R9 ( Ppatg18 ), Ppvps15 ( Ppvps15), WDY53 ( Ppvac8), and SMD1163 ( pep4, prb1) cells were grown in minimal medium containing 14C-valine for 18 h. The cells were pellete d and resuspended in medium lacking amino acids and nitrogen and containing 10 mM valine. The production of TCA-soluble radioactivity was measured at 2, 5, 8, and 24 h of chase and the rates calculated by linear regression of the slope of the line. The rates represent the mean S.E. of 5-7 trials. 89

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Figure 3-2. Pexophagy is blocked at an early sequestration event in cells lacking PpAtg9. Wild type cells, GS115 (A, C) and WDY70 (B, D) and cells lacking PpAtg9, R19 (E, G, I, K) and WDY71 (F, H, J, L) were grown in YNM for 24 -36 h. At that time, cells were switched to medium containing 2% glucose (A, B, E, F, I, J) or 0.5% ethanol (C, D, G, H, K, L) for 3 h. Cells were fixed with potassium permanganate and prepared for viewing on a JEOL 100CX transmission electron microscope (A C, E, G, I, K) or viewed in situ by fluorescence microscopy (B, D, F, H, J, L). When GS115 and WDY70 cells were adapted to glucose for 3 h, many of the cells lacked peroxisomes while in others the vacuole was found to virtually surround the peroxisome cluster (A, B). In R19 and WDY71 cells, peroxisome clusters were evident in virtually all the cells. The vacuole was either oblong with a cup-like de pression at the surface adjacent to the peroxisomes (E, F) or round with short arm-like segments (I, J). By fluorescence microscopy, the vacuole was observed by staining with FM 4-64, the peroxisomes by expressing BFP-SKL, and the MIPA and pexophagosomes by expressing GFP-PpAtg8. GFP-PpAtg8 was observed at the MIPA in WDY70 cel ls (B, arrow), but appeared as foci in WDY71 cells lacking PpAtg9 (F and J, arrowheads). During etha nol adaptation, individual peroxisomes were sequestered into pexophagosomes (C, arrow) that contained PpAtg8 (D, arrow). No pexophagosomes were evident in R19 and WDY71 cel ls lacking PpAtg9 (G, H, K, L). In the absence of macropexophagy, PpAtg8 re sided in numerous foci (H and L, arrowheads). N, nucleus; P, peroxisome; V, vacuole. 90

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Figure 3-3. Cellular localization of PpAtg9. TC14 expressing PpAtg9-GFP behind the endogenous PpAtg9 promoter (A, B), AJM27 expressing GFP-PpAtg9 behind the GAPDH promoter and dsRFP-HDEL (C), AJM44 expre ssing mRFP-Atg9 behind the GAPDH promoter and CoxIV-GFP (D), AJM19 expressing mR FP-PpAtg9 behind the GAPDH promoter and Sec13-GFP (E), and AJM18 expressing mRFPPpAtg9 behind the GAPDH promoter and Sec7GFP (F) were grown in YPD and the cellular di stribution of the GFP and mRFP tagged proteins visualized by in situ fluorescence microscopy. In TC14 cells, PpAtg9-GFP was present in foci situated at the cell periphery (arrows). The GFPPpAtg9 structures appeared to be juxtaposed to the endoplasmic reticulum containing dsRFPHDEL (C) and mitochondria containing CoxIVGRP (D), but distinct from intermediate vesicl es containing Sec13-GFP (D) and Golgi apparatus containing Sec7-GFP (E). 91

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Figure 3-4. PpAtg9 relocates from peripheral structures to th e sequestering membranes during glucose-induced pexophagy. TC3 cells expres sing both BFP-SKL behind the AOX1 promoter and GFP-PpAtg9 behind the GAPDH promoter were grown in YNM medium in the presence of FM 4-64, adapted to YND medium for 0 h (Panel A) and 3 h (Panel B), and the cellular distribution of GFPPpAtg9 visualized by in situ fluorescence microscopy. Peroxisomes were identified by the presence of BFP, which was targeted by its SKL signal and the vacuole by the 92

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red dye FM 4-64. At 0 h, many cells had B FP-containing peroxisomes and a single round vacuole. GFP-PpAtg9 was localized to foci near the cell periphery (arrowheads). At 3 h of glucose adaptation, numerous profiles of vacuol es with arm-like exte nsions surrounding the BFP-containing peroxisomes could be observed. GFP-PpAtg9 was present at perivacuolar structures (arrows) and at the vacuole membrane (double arrowhead). In panel C, images from 0-2 h of glucose adaptation depict the moveme nts of GFP-PpAtg9 from peripheral structures (arrowheads) to the perivacuolar structures (arrows) and sequestering and vacuole membranes (double arrowhead). In panel D, ASK2 ce lls expressing HA-PpAtg9 behind the endogenous PpAtg9 promoter were grown in methanol medium and then adapted to glucose medium. At 0-8 h, equivalent numbers of cells based on the optical density (OD600) of the cultures were solubilized in SDS, and the pr oteins separated by SDS-PAGE. HA-PpAtg9 was then visualized by Western blotting using anti-HA antibodies and quantified using a Typhoon laser scanner. The values were normalized to 0 h and presented as the mean SE (n = 6). 93

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Figure 3-5. Trafficking of PpAt g9 in mutants defective in earl y sequestration events. TC6 ( Ppatg11), TC9 ( Ppatg18), and TC11 ( Ppvps15) mutants expressing GFP-PpAtg9 behind the GAPDH promoter were grown in YNM medium in the presence of FM 4-64 then adapted to YND medium for 3 h and the cellular dist ribution of GFP-PpAtg9 visualized by in situ fluorescence microscopy. The vacuoles in the Ppvps15, Ppatg11 and Ppatg18 mutants were either round, flattened, or slightly indented. GFP-PpAtg9 was present in peripheral structures (arrowheads) in the Ppvps15 and Ppatg11 mutants. However, in Ppatg18 mutants, GFPPpAtg9 was visualized at the large sometimes diff use perivacuolar struct ures (arrows) and the vacuole membrane (double arrowheads). 94

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Figure 3-6. Trafficking of PpAt g9 in mutants defective in intermediate sequestration events. TC19 ( Ppatg2) and TC5 (Ppatg7) mutants expressing G FP-PpAtg9 behind the GAPDH promoter were grown in YNM medium in th e presence of FM 4-64 then adapted to YND medium for 3 h and the cellular distri bution of GFP-PpAtg9 visualized by in situ fluorescence microscopy. The vacuoles in Ppatg2 and Ppatg7 mutants were indented with an occasional short segmented arm-like extension. In these mutants, GFP-PpAtg9 localized to peripheral structures (arrowheads) and the PVS at the vacu ole surface (arrows), but was absent from the vacuolar membrane. Similar results were obtained in TC8 ( Ppatg2) and TC4 ( Ppatg7) cells. 95

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Figure 3-7. Trafficking of Pp Atg9 in mutants defective in late sequestration events. TC7 ( Ppatg1) and TC12 ( Ppvac8) mutants expressing GFP-PpAtg9 behind the GAPDH promoter were grown in YNM medium in the presence of FM 4-64 then adapted to YND medium for 3 h and the cellular distribution of GFP-PpAtg9 visualized by in situ fluorescence microscopy. The vacuoles in Ppatg1 and Ppvac8 mutants were indented with multiple segmented arm-like extensions that surrounded the peroxisome clus ter. GFP-PpAtg9 was visu alized at the vacuole membrane but appeared to be more concentrated at the sequestered membrane extensions of the vacuole (double arrowheads). In addition, GFP-PpAtg9 was found at the PVS situated at sites of the vacuole surface adjacent to the arm-like extensions (arrows). 96

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Figure 3-8. PpAtg9 domains required for function and trafficking. In panel A, wild type GS115, R19 ( Ppatg9), and R19 cells expressing mR FP-PpAtg9, GFP-PpAtg9, GFP-Ppatg9 N ( M1Y221), or GFP-Ppatg9 L3 ( T471 A523) were grown in YNM medium for 36 h. All proteins were expressed behind the GAPDH promoter. Ce lls were then switched to glucose medium. Aliquots were removed at 0 and 6 h of adapta tion, the cells lysed, and AOX activities measured as described under Experimental Procedures. Th e data are expressed as a percentage of AOX remaining at 6 h relative to 0 h and represent the mean S.E. of 4-6 trials. ASK5 (B) and ASK4 (C) cells were grown in YNM for 20 h in th e presence of FM 4-64. The cells were then transferred to YND medium for 3 h and then visualized by fluorescence microscopy. GFPPpatg9 N was found at the Atg9-PC (arrowheads), PVS (arrows) and the vacuolar membranes. GFP-Ppatg9 L3 was absent from the Atg9-PC and PVS, but found in a compartment that resembled the endoplasmic reticulum (large arrows). 97

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Figure 3-9. Perivacuolar stru ctures contain PpAtg9 and Pp Atg11. AJM33 expressing GFPPpAtg18 and mRFP-PpAtg9, AJM25 expressing GFP-PpAtg2 and mRFP-PpAtg9 and AJM32 expressing GFP-PpAtg11 and mR FP-PpAtg9 were grown in YNM medium then adapted to YND medium for 2 h and the cellula r distribution of GFP and RFP tagged proteins visualized by in situ fluorescence microscopy. PpAtg9 was found at peripheral structures (arrowheads) and PVS (arrows) and within the vacuole delineated by PpAtg18 and PpAtg11. The Atg9-PC did not contain PpAtg2, PpAtg11 or PpAtg18. GFP-PpAtg2 resided in distin ct vesicles of which a few were in close association with the PVS. M eanwhile, virtually all th e PVS contained both GFPPpAtg11 and mRFP-PpAtg9 (arrows). 98

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Figure 3-10. PpAtg9 does not colocalize with PpAtg8 structures. WD Y75 expressing GFPPpAtg8 and mRFP-PpAtg9 were grow n in YNM then adapted to glucose or ethanol medium for 2 h and the cellular distribution of GFP and RFP tagged protei ns visualized by in situ fluorescence microscopy. During glucose adapta tion (panel A), mRFP-Pp Atg9 was found within the vacuole and at the PVS (large arrows), but ab sent from PpAtg8 foci and MIPA (arrowheads). When cells were transferred from methanol to ethanol medium (pan el B), mRFP-PpAtg9 was present within the vacuole and at the peripheral compartment (small arrow) and the PVS (large arrow), but not at the PpAtg8 foci or pexophagosome (arrowheads). 99

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100

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Figure 3-11. Model of PpAtg9 trafficking dur ing pexophagy. During growth in YNM, PpAtg9 (green) is in peripheral compar tment (Atg9-PC), PpAtg11 (red) at the vacuole membrane, and PpAtg2 (brown) cytosolic. Upon glucose-induced pexophagy, PpAtg9 is trafficked from the Atg9-PC, to the PVS juxtaposed to the vac uole which also contains PpAtg11, and to the sequestering membranes (SM) that surround the per oxisomes (blue). Also at this time, PpAtg2 becomes associated with unknown structures situated near the PVS. The trafficking of PpAtg9 from the Atg9-PC to the PVS requires PpAtg11 and PpVps15. The movements of PpAtg9 from the PVS to the vacuolar and sequestering membranes require PpAtg7 and PpAtg2. The formation of the sequestering membranes from the vacuole requires PpAtg18, but the movements of PpAtg9 to the vacuolar membrane proceeds normally in the absence of this protein. Finally, the fusion of th e sequestering membranes that enable the incorporation of the peroxisomes into the vacuole is guided by the micropexophagy-specific membrane apparatus (MIPA) and requires PpVac8 and PpAtg1. 101

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CHAPTER 4 PPATG9 TRAFFICKING DURING MICROPEXOPHAGY IN PICHIA PASTORIS5 PpAtg9 encodes a 102 kDa membrane protein with five or more putative membrane spanning domains. Atg9 is structurally conserved throughout eukaryotes with each containing the APG9 domain (Pfam ID, PF04109). Unlike lowe r eukaryotes, there appears to have evolved two homologues, Atg9L1 and Atg9L2, in vertebra tes including fish, amphibians, birds, and mammals.(Yamada et al. 2005) Interestingly, both forms are functional for autophagosome formation.(Yamada et al., 2005) The insertion of Atg9 into the lipid bilayer occurs independent of a recognizable signal peptide. Furthermore, placing GFP at the Nor C-terminus of PpAtg9 has no effect on its function. We have recently described the trafficki ng of Atg9 during glucoseinduced micropexophagy in Pichia pastoris.(Chang et al., 2005) Here we will summarize our findings and describe those cellular events responsible for PpAtg9 trafficking. During exponential growth, we have shown that PpAtg9 resides in peripheral vesicles lacking Sec7 or Sec13 but that sometimes colo calizes with markers of the rough endoplasmic reticulum (dsRFP-HDEL) and the mitochondria (CoxIV-GFP).(Chang et al., 2005) A majority of the ScAtg9 localize with mitochondria or in vesicles situated near mitochondria.(Reggiori et al. 2005) However, these authors have also shown that only a small fraction of the PpAtg9 localize with the mitochondria.(Reggiori et al. 2005) The function of Atg9 at the mitochondria remains unclear, but it may have a role in th e selective degradation of damaged mitochondria (e.g., mitophagy). Because of the unique characteristics of the PpAtg9 vesicles, we have classified them as the Atg9 peripheral compartm ent (Atg9-PC) which likely arises from the endoplasmic reticulum (ER). In fact, we have identified a putative ER exit motif, which upon 5 Article reprinted with permission from the publishe r: Schroder, L.A. and W.A. Dunn, Jr. (2006) PpATG9 trafficking during micropexophagy in Pichia pastoris. Autophagy. 2(1):52-54. 102

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deletion (Atg9L3) results in the accumulation of PpAtg9 within the ER.(Chang et al. 2005) This di-acidic ER exit motif is positioned within a highly conserved regi on of Atg9 situated between putative transmembrane domains in P. pastoris S. cerevisiae, D. melanogaster C. elegans, and H. sapiens (Fig. 1). This di-acidic (YxDxE) motif was first identified at the Cterminal tail of VSV-G protein and later shown to be functional in other membrane proteins (Barlowe, 2003). Furthermore, a di-acidic YKDAD motif has been sh own to be essential for the exit of the cystic fibrosis transmembrane c onductance regulator (CFTR) from the ER.(Wang et al. 2004) In the latter case, this motif is located not at the C-te rminus, but at a surface-exposed loop as is likely the case for Atg9. The YKDAD motif is similar to the FNELE sequences in CeAtg9 and HsAtg9L1 and to the FNELD sequence present in DmAtg9. Th e di-acidic ER exit motifs in PpAtg9 and ScAtg9 have similarities to motifs found in both CFTR and Kir2.1, a potassium channel protein (Fig. 4-1). ScAtg9 has been shown to cycle between a peripheral compartment (e.g., mitochondria) and the preautophagosomal structure (PAS).(Reggiori et al. 2004; Reggiori et al. 2005) We suggest that Atg9 transits the Atg9-PC during this cycling (Fig. 2). However, the dynamics of the Atg9-PC and its relationship with th e mitochondria and PAS have not yet been defined. Upon the onset of glucose-induced micrope xophagy, a number of even ts occur prior to the formation of the sequestering membranes (S M), a portion of the vacuole membrane, that surround the peroxisomes (Fig. 4-2). First, PpAt g11 transits from a single site at the vacuole surface presumed to be the PAS to multiple peri vacuolar structures (PVS) (Step 1, Fig. 4-2). Next, PpAtg9 is transferred from the Atg9-PC to the PVS (Step 2, Fig. 4-2). The PVS is unique from the PAS in that it does not contain PpAtg8 a nd is situated at multiple foci at the vacuole surface where it appears to direct the formation of the SM in much the same way that the PAS 103

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organizes the formation of the autophagosome. The trafficking of PpAtg9 to the PVS requires PpAtg11 and PpVps15. It remains unclear whether PpAtg9 transits from the PAS to the PVS or arrives directly from the Atg9-PC. At this time, cytosolic PpAtg2 becomes associated with peripheral foci and the PVS, while cyto solic PpAtg18 becomes bound to the vacuolar membrane.(Guan et al., 2001; Stromhaug et al., 2001) The PpAtg2 foci do not contain PpAtg9, but are distinct from the Atg9-PC. However, th e assembly of this Atg2 compartment (Atg2-C) requires PpAtg9 suggesting a transi ent association. PpAtg9 is then transferred from the PVS to the vacuole membrane and newly forming SM (Ste p 3, Fig. 4-2). PpAtg2 is required for PpAtg9 trafficking from the PVS to the SM, while PpAtg18 is essential for the formation of the SM. The assembly of the SM is best described by a comb ination of incomplete and complete septation from the vacuole followed by membrane fusion events to form arm-like structures. Since PpAtg11 is unique to pexophagy and found at the SM, we propose this protein may act to recognize and guide the SM around the peroxisomes. The SM fusion events require PpAtg24 and PpVac8.(Ano et al., 2005; Chang et al., 2005; Wohlgemuth et al. 2007) As the SM is being formed, the PAS assembles a unique micrope xophagy-specific membrane apparatus (MIPA) which contains PpAtg8 and PpAtg26 (Step 4, Fig. 4-2). In addition to those enzymes (e.g., PpAtg3, PpAtg4, and PpAtg7) essential for the conjugation of PpAtg8 to phosphatidylethanolamine (Oku et al. 2003; Mukaiyama et al., 2004), the formation of the MIPA requires PpAtg11 (unpublis hed data) and PpAtg9.(Chang et al., 2005) Although neither PpAtg11 nor PpAtg9 are associated with the MIPA, their role in its formation suggests these proteins may be at least transi ently associated with the PAS. Once the MIPA is assembled it directs the final membrane fusion of the SM th ereby incorporating the peroxisomes into the vacuole for degradation (Step 5, Fig. 4-2). We suggest that glucose-induced micropexophagy 104

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proceeds through a complex but well-organized se quence of molecular events to ensure the selective and efficient degradation of superfluous peroxisomes. Figure 4-1. Alignment of the putative ER exit motifs of Atg9. Th e amino acid sequence between putative transmembrane (TM) domains of Atg9 from P. pastoris S. cerevisiae and H. sapiens were aligned with structural homologs from D. melanogaster and C. elegans. Amino acid identities (--) and gaps () are indicated and the putative ER exit domai n aligned with similar motifs in CFTR and Kir2.1. 105

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Figure 4-2. Trafficking of PpAtg9 and other Atg proteins during micropexophagy. When P. pastoris are grown in methanol medium, PpAtg proteins can be found in a number of compartments (outside the box): endoplasmic reticulum (ER), Atg9 peripheral compartment (Atg9-PC), pre-autophagosomal st ructure (PAS), mitochondria, cy tosol and the vacuole. Upon adaptation to glucose medium and induction of micropexophagy, many of the PpAtg proteins traffic to compartments unique to micropexopha gy (inside the box): Atg2 compartment (Atg2C), perivacuolar structures (PVS), sequester ing membranes (SM), and micropexophagy-specific membrane apparatus (MIPA). The trafficki ng of PpAtg9 and other Atg proteins during micropexophagy is summarized in this di agram and discussed in the text. 106

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CHAPTER 5 TARGETING MOTIFS OF ATG9 Introduction Protein degradation keeps the cell in constant homeostasis even during times of starvation and stress. Autophagy is the foremost mechanism by which cellular milieu, from proteins to organelles, is delivered in a hi ghly regulated fashion to the lytic compartmentthe lysosome or the vacuolefor degradation. Multiple select ive and nonselective modes of autophagy exist in yeast, namely autophagy, pexophagy, and mitophagy (Wang and Klionsky, 2003; Dunn et al., 2005). Autophagy is turned on when cells are starved for amino acids. Pexophagy occurs in methylotropic yeasts during adaptation from metha nol to ethanol or glucose as peroxisomes are degraded with the media shift (Dunn et al., 2005). When stationary cu ltures of S. cerevisiae are switched to medium containing lactose, mito chondria are degraded by mitophagy (Abeliovich, 2007). Pichia pastoris is a methylotropic yeast able to synthesize the enzymes to consume methanol as sole dietetic source. The peroxisomes are synthesized when P. pastoris is exposed to methanol to metabolize this carbon source. When the cells are adapted to glucose the peroxisomes are rapidly sequestered and degraded by the va cuole via key micropexophagic eventsthe superfluous organelle is no longer needed and is cons umed and degraded to provide the building blocks for protein synthesis (Dunn et al. 2005). These events include the recognition of the peroxisome by the vacuole a nd the invagination of the vacuole with the protrusion of vacuolar arms that surround then engulf the collection of peroxisomes as a whole for degradation. If P. pastoris is adapted from methanol to et hanol (rather than glucose), the peroxisomes are selectively sequestered indivi dually by pexophagosomes that fuse with the vacuole. 107

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The sequestration events of glucose-induced micropexophagy can be phenotypically characterized as the sequentia l events nucleation, expansi on, completion and degradation. Nucleation of the sequestering membranes oc curs with the budding of the sequestering membranes (SM) from the vacuole and requi res Atg18. Expansion of the sequestering membranes to surround the peroxisomes follows from perivacuolar struct ures (PVS), involving Atg2, Atg11 and Atg9. To assist in tetheri ng of the SM, the micropexophagic-specific membrane apparatus (MIPA) forms betw een the expanding sequestering membranes (Mukaiyama et al., 2004; Ano et al., 2005). The assembly of th e MIPA requires Atg7 and Atg8. Completion is the final enveloping of the peroxisomes by the SM, with the participati on of Vac8. Atg9 is a transmembrane protein required for SM expansion during pexophagy and autophagy (Chang et al., 2005). That is, in the absence of At g9 the SM only partially engulfs the peroxisomes. Upon ER synthesis, Atg9 transits to the Atg9 peripheral compartment (PC9). When cells adapt to glucose, Atg9 moves to the perivacuolar structure (PVS) and to the preautophagosome structures (PAS). From the PVS, Atg9 transits to the sequestering membranes (SM) (Schroder and Dunn, 2006). The PV S is present at the base of the SM while the PAS is localized adjacent to the tips of the SM (Chang et al. 2005). Atg11 recruits Atg9 to the PVS which has been shown to contain Atg9 and Atg11 but not Atg2 in P. pastoris Atg9 is required for early and late seque stration; more specifically, Atg9 is required for MIPA formation during late sequestration. Recent data in Saccharomyces cerevisae indicate a direct interaction between ScAtg9 and ScAtg11 at the PAS (Chang and Huang, 2007; Mari and Reggiori, 2007). The interaction between ScAtg11 and ScAtg9 appear s to require the first two coiled-coil regions of ScAtg11 as well as amino acids 154-200 of th e N-terminus of ScAtg9 but not the last 200 108

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residues of the C-terminus of ScAtg9 (Chang and Huang, 2007). Interestingl y, actin is essential for the trafficking of ScAtg9 and ScAtg11 to the PAS (Reggiori et al. 2003; Yen et al. 2007). In order to better clarify its dynamic role of Atg9 in pexophagy, we have mutated conserved regions of Atg9. De letion of the N-terminus resu lted in a non-functional albeit properly sorted mutant phenotype that localizes to the vacuole and SM. There are two ER exit domains within Atg9. Deletion of the region between the third and fourth transmembrane domains, an Atg9 loopmost importantly am ino acids E491 and E494suggests that this domain is critical for ER exit. Deletion of th e region past the last tr ansmembrane domain also resulted in ER localization. The PC9 exit domain encompasses amino acids QQHKA at the extreme C-terminal end of Atg9. Localization to the SM requires amino acids W607 and Y611. All the mutants presented exhibi t altered trafficking save atg9 N which localizes to the SM along the vacuole but does not function. Ind eed, none of the atg9 mutants was functional, revealing that Atg9 trafficking to the SM is essential for micropexophagy. Materials and Methods Yeast Strains And Media Table 1 lists the yeast strain s utilized in this study. Pichia pastoris was routinely cultured and incubated at 30C in YPD (1% Bacto yeast extract, 2% Bacto peptone, and 2% dextrose). To induce peroxisome proliferation, P. pastoris was grown in YNM (0.67 % yeast nitrogen base, 0.4 mg/l biotin, and 0.5% methanol ); to induce peroxisome abi ogenesis, cells grown in YNM were transferred to YND (0.67 % yeast nitrogen base, 0.4 mg/l bi otin, and 2% glucose) or YNE (0.67 % yeast nitrogen base, 0.4 mg/l biotin, and 0.5% ethanol). For induction of macroautophagy, yeast was grown in nitrogen starvation medium (SD-N) containing 0.17 % yeast nitrogen base (without NH4SO4 and amino acids except for auxotrophic amino acids as needed) and 2% glucose. For plates, all me dia included 2% agar. Auxotrophic strains had 109

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arginine or histidine or both added at 40 g/l as necessary. Vectors were amplified in the DH5 strain of E. coli and cultured with ampicillin (100 g/ml) in LB broth (0.5% Bacto yeast extract, 1% Bacto tryptone with 1% NaCl) at 37 C. 25 g/mL of Zeocin was added to DH5 and 100 g/ml of Zeocin was added to P. pastoris Yeast Transformation Once yeast cultures reached saturation density ( A600 = 1.0), they were harvested and treated with 10 mM DTT in YPD containing 25 mM HEPES, pH 8, for 15 min, all at 30C. Afterwards, the cells were washed twice with cold water, wa shed once with 1 M sorbitol, and resuspended in 1 M sorbitol. Electroporation of linearized vector (40 l of yeast cells to 0.2-1 g of linearized vector) occurred in a 0.2 cm ga p cuvette (Bio-Rad, Hercules, CA) with electroporation conditions as follows: 1.5 kV, 25 F, 400 (Gene Pulser, Bio-Rad Corp.). Histidine was the remaining auxotrophy for Atg9; cells transformed with vectors containing the His4 gene were screened on plates containing 0.67% yeast nitrogen base without amino acids, 2% glucose, 1M sorbitol, 0.4 mg/l biotin, and 2% agar and checked for colonies after 3-5 d at 30 C. The other screening method utilized was the incorporation of the zeo cin resistance gene (zeoR) and cells transformed with (zeoR) vectors were grown on YPD with 100 g/ml Zeocin. Isolation Of Gsa Mutants And Cloning Of GSA Genes By Restriction Enzyme Mediated Integration (REMI) Mutagenesis Mutagenesis by random insertion the pREMI-Z vector (provided by Dr. Ben Glick, Univ. of Chicago) for genomic incorporation of the Zeocin resistance gene into the genome of P. pastoris has been described previously (Stromhaug et al., 2001). Direct colony assays verified gsa mutants as described before(Stromhaug et al. 2001). The site of insertion of the pREMI-Z and identification of the disrupted has been explained (Stromhaug et al., 2001). The GSA14 gene (NCBI accession number AY075105) was assembled a nd is homologous to to following: ScAtg9 110

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of Saccharomyces cerevisiae NP_596247 of Schizosaccharomyces pombe NP_180684 of Arabidopsis thalian XP_331198 of Neurospora crass NP_611114 of Drosophila melanogaster NP_503178 of Caenorhabditis elegans, and NP_076990 of Homo sapiens (Yamada et al., 2005). Homologous residues 190-679 contain a minimum of 5 putative transmembrane domains. The pREMI-Z disruption of PpAtg9 occurred after th e first putative transmembrane domain at aspartic acid 245. The null mutant of Atg9 was made as follows. ATG9 was cloned and inserted into the pUC19 vector. ScAR G4 was amplified from genomic DNA by PCR using a forward primer 5-CCACGTTCTAGAGGTAGATGT-3 that contai ned an XbaI site and a reverse primer 5-GATGAAGTCCGCGGAGTACCA -3 that contai ned a SacII site. ScARG4 with its endogenous promoter was ligated within the ATG9 gene that had been cut with SacII and XbaI thereby replacing the en tire coding region of ATG9 The product was then transformed in PPF1 cells and the resulting mutants screen by direct colony assays. WDY55 was constructed by mating R19 with PPF1 and screen ing the haploids for double hisand argauxotrophy. Those double auxotrophs defective in pexophayg were then identified by dire ct colony assays. WDY100 cells expressing were constructe d by transforming PPF1 cells with pWD26 (PGAPDH RFP-Atg17, ARG4). Construction of Ppatg9 Expression Vectors The GFP-Atg9 and RFP-Atg9 expression vect ors (pTC1 and pAJM6) as well as the pGFP-atg9 N and pGFP-atg9 L3 have been described previously (Chang et al. 2005). The GFP-atg9 QQHKA mutant was made as follows. Pp ATG9 minus the QQHKA sequence was amplified from pTC1 with forward primer 298 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 that co ntained the KpnI restriction site and reverse primer 5-CAAATTATGCCT CGA GCTGTTAACTAA-3. The ATG9 sequence minus 111

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the last 5 amino acids was then cut with KpnI and XhoI. GFP was excised from pIB2 from the KpnI and the XhoI rest riction sites. The Pp atg9 QQHKA was ligated to GFP at the KpnI and XhoI restriction sites, amplified, verified by se quencing, and the vector used to transform R19 ( his4 atg9 :: zeoR) cells. The GFP-atg9 579-668 mutant was composed as follows. First, the coding region between the endogenous promoter upstream of the N-terminus and amino acid 579 of Pp ATG9 was amplified from pTC1 with forward primer 298 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 that co ntained the KpnI restriction site and the reverse primer 5-CAGGAATCGA AAG CTTGCAAATAGT-3 that contained the HindIII restriction site. Next, the region betw een amino acid 669 and the C-terminus of Pp ATG9 was amplified from pTC1 with fo rward primer 5-GTGTTCATGTTGAT AAGC T T GGATATGTA3 that contained the HindIII restrict ion site and reverse primer 266 5CAAGATCTATGCTCGAGAACAAAT AATGCCTTATG CTGTTGACTAA-3 that contained the XhoI site. The parent vector pPS55 was cut with KpnI and XhoI. The N-terminus of Atg9 including the promoter region through amino acid 579 region a nd the C-terminus of Atg9 not including the stop codon and pPS 55 with GFP upstream of the XhoI site were ligated together, amplified, verified by sequencing, and th e vector used to transform R19 ( his4 atg9:: zeoR) cells. The GFP-atg9 489-502 mutant was constructed as follows The region between the N-terminus and amino acid 489 of ATG9 was amplified from pTC1 with forward primer 298 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 with the KpnI restriction site and with reverse primer 5-AATGCGATCCACCTTCGA T TCTCTTAAATTAC-3 with the HindIII restriction site. The region between amino acid 502 and the C-terminus of ATG9 gene was amplified from pTC1 with fo rward primer 5-CATTTTCAACAGAA A GCT T AACATTTCCA3 with the HindIII restriction si te and with reverse primer 5112

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CATTATTATTCAAGA C C GCG GAATGAAAA-3 with the SacII re striction site. The digest products were ligated together, amplified, the vector verified by sequencing, and used to transform R19 ( his4 atg9 :: zeoR) cells. The GFP-atg9(W607A/Y611A) mutant was generated as follows. ATG9 gene upstream of the N-terminus and flanking amino acids 607-611 was amplified by PCR from genomic DNA with the forward primer 298 5-GCAGGCTAGGGT ACCGGTACTGGCACATT-3 and with the reverse primer 5-CTTCCTCTGTGTGGGCTTTCCCTTCCGCTTCTTGAGGAA-3 with a KpnI restriction site. At the same time, At g9 flanking amino acids 607-611 and through the stop codon at the C-terminus was amplified from genomic DNA with the forward primer 5CTTCCTCAAGAAGCGGAAGGGAAA GCCCACACAGAGGAA-3 and with the reverse primer 5-GTTTTGGACTCGAGGGTACTAATGCTTCA TT-3. The product sizes were first verified by restriction digest, then amplified together wi th the forward primer 298 5GCAGGCTAGGGTACCGGTACTGGCACATT-3 and with the reverse primer 5GTTTTGGACTCGAGGGTACTAATGC TTCATT-3. The size of the amplified product atg9(W607A/Y611A) was verified by restriction digest and by sequencing and afterwards digested with the restriction enzymes KpnI and X hoI. The product of the re striction digests from atg9(W607A/Y611A) was inserted into the KpnI and X hoI sites of pTC1 and the vector used to transform R19 ( his4 atg9 :: zeoR) cells. Qualitative And Quantitative Assessment Of Alcohol Oxidas e (AOX) Degradation The direct colony assay was us ed to verify and select atg9 deletion mutant colonies, as described in previous publica tions (Tuttle and Dunn, 1995; Yuan et al., 1997). For clarification, colonies were replica-plated from Zeocin to YNMH plates and incubated for 3-4 days. Replicas on nitrocellulose were then placed on YNDH plat es for 12-15 h. The replica nitrocellulose was dipped in liquid nitrogen with a tw eezer for 20 seconds with care not to sheer the nitrocellulose 113

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to lyse the cells. Placing the nitrocellulose on Whatman paper soaked in 33 mM potassium phosphate buffer, pH 7.5, containing 3.4 U/ml horseradish peroxidase 0.53 mg/ml 2,2'-azinobis(3-ethylbenz-thazoline-6-sulf onic acid), and 0.13% methanol at room temperature for 60-90 minutes allows visualization of colonies that ar e unable to degrade AOX. For the liquid medium assay, cells are grown to satura tion in 20 mls YNM with methanol as carbon source. After 40 hours incubation, 0.4 g of powdered glucose or 100 l of ethanol was added. At 0 and 6 hours of glucose or ethanol adaptation (8.0 OD600), 2 ml aliquots of glucos e or ethanol were pelleted and resuspended in 1 ml 20 mM Tris, pH 7.5, containing 50 mM Na Cl, 1 mM EDTA, 1 mM PMSF, 1 g/ml pepstatin A, and 0.5 g/ml leupeptin. 0.5 ml glass beads (425-600 microns) were used to lyse the cells by vortexing. The glass beads and cellular debris were removed by centrifugation and AOX measured by adding 50 l of this extract to 3 mls of reaction mix containing 3.4 U/ml horseradish peroxidase and 0.53 mg/ml 2,2'-azino-bis(3-ethylbenzthazoline-6-sulfonic acid) in 33 mM potassium phos phate buffer, pH 7.5 (Tuttle et al. 1993; Tuttle and Dunn, 1995). The addition of 10 l me thanol initiated the reaction, which was then continued at room temperature for 15 -30 minutes. The assay was quenched by addition of 200 l of 4 N HCl and the absorbance read at 410 nm. Western Blot Analysis Cells were grown for 16 hours in 20 ml YNM and then adapted to YND or YNE. Cell aliquots were collected by centrifugation. The cells were pellete d and lysed in 100-150 L of 67 mM Tris, pH 6.8, 2% SDS, 10% glycerol, 0.1 % bromophenol blue, 1.5% DTT solution and 2 L of PIC (200 mM phenylmethylsulfonyl fluor ide (PMSF), 14.5 mM pepstatin A, 105 mM leupeptin in DMSO) by vortexing (four times at one -minute intervals) with glass beads for SDSPAGE. After the sample preparat ion, they were heated at 100 C for 5 minutes and cell debris 114

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was removed by centrifugation. Fifteen to thirty microliters of each sample were loaded onto 10% or 10-15% SDS-PAGE. The proteins were then transferred to nitrocellulose by Trans-Blot SD-Dry Transfer Cell (Bio-Rad Laborator ies, Hercules, CA) for 1.15-1.5 h, depending on whether the proteins being transf erred were small or large respec tively. The blots were blocked in 5% nonfat dried milk in PBS containi ng 0.1% Tween 20 (PBS-T) for 2 hours at room temperature, incubated with mouse anti-GFP or mouse anti-HA primary antibody (Covance Inc., Princeton, NJ) overnight at 4C, and then incuba ted in 2% nonfat dried milk with secondary goat anti-mouse antibody conjugated with HRP (Cov ance Inc., Princeton, NJ) for 2 h at room temperature. After each step the blots were wash ed 3 times in PBS-T. HRP was detected using ECL-plus (Amersham, Piscataway, NJ) and quantified using the Typhoon 9400 laser scanner (Molecular Dynamics, Sunnyvale, CA). Measurements of Protein Degradation The degradation of cellular proteins during nitr ogen starvation was performed as described previously (Tuttle and Dunn, 1995; Yuan et al., 1999). Endogenous proteins were radiolabeled with 1 Ci/ml 14C-valine for 16 h in 0.67 % yeast nitrogen base, 2% glucose, 0.4 mg/l biotin, and 40 g/l histidine or arginine (if need ed). The cells were then washed and switched to nitrogen starvation medium containing 0.17 % yeast nitrogen base (without amino acids and NH4SO4) and 2% glucose and supplemented with 10 mM valin e. Aliquots were removed at 2 24 hours of chase and trichloroacetic acid (TCA ) added to a final concentrati on of 20%. Acid soluble and insoluble radioactiv ity was separated by centrifugation and the radio activity present measured by scintillation counting. The rates of protein degradati on were calculated from the slopes of the linear plots of TCA-sol uble radioactivity versus time of chase. 115

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Fluorescence Microscopy and FM 4-64 Labeling Cells were grown in either YPD or YNM fo r 20 hours for visualiza tion of GFP or RFP fusion protein expression. Cells grown on YNM medium were then transferred to YND or YNE for 2 hours. FM 4-64 (Molecular Probes, Eugene, OR) was added to a final concentration of 20 g/ml and the cells incubated for 2-12 hours prior to nutrient adaptation. The cells were washed of unbound FM 4-64 and examined immediately using a Zeiss Axiophot fluorescence microscope. Image capture was done using a SP OT camera (Diagnostics Instruments, Inc., Sterling Heights, MI) interfaced with IP Lab software. Results Atg9 and Atg17 Occasionally Co-localize Atg9 is known to localize transiently to the PAS in S. cerevisiae (Reggiori et al., 2005). We have previously shown that Atg9 did not co localize with Atg8, but studies have shown that these proteins may transiently interact (Te llkamp, unpublished observations). To determine whether Atg9 is present in a compartment dis tinct from the PAS, monomeric RFP tagged Atg9 was co-expressed with GFP-Atg17, a marker for the PAS (Oku et al., 2006). mRFP-Atg9 did not completely co-localize with the GFP-Atg17 (Figure 5-1). The GFP-Atg17 localized to a number of dots not containing mRFP-Atg9, but remarkably near dots containing mRFP-Atg9. Meanwhile, GFP-Atg17 did localiz e with mRFP-Atg9 at the PC 9 and PVS. Based on our observations, we propose that th e Atg9-containing PVS abuts the Atg17-containing PAS, and that Atg9 may transit to the PAS during glucose-induced micropexophagy. Ppatg9 Partial Deletion And Site-Specific Mutants Are Defective In Glucose-Induced Pexophagy And Starvation-Induced Autophagy When the available carbon source is restricted to methanol, Pichia pastoris responds to this environmental stress by synthesizing peroxisomal enzymes such as alcohol oxidase (AOX) 116

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for degradation. When this stress is relieved by a media switch from methanol to glucose, the peroxisomes are selectively sequestered and de graded via micropexophagy. A number of ATG gene products have been shown to be required for autophagic induction. We have shown that Atg9 transports from unique peri pheral vesicles to the expanding SM where it functions. Our aim was to delete highly conserve d regions of Atg9 or sequences identified by motif databases to be important. Thus, the following regions were deleted: the N-terminus prior to the first transmembrane domain (atg9 N), the loop between the third a nd fourth transmembrane domains (atg9 L3), the C-terminal region just past the last transmembrane domain (atg9 579-668), a motif present in Pex5 and Pex14 (atg9-W607A/ Y611A) and recognized by Pex13 and Pex14 for Pex5 binding [the significance of the domain present in Pex14 is not understood], and a motif with high homology to the ER export mo tif for transmembrane proteins (atg9 QQHKA 881885) (Figure 5-2). Smaller motifs of higher homology of the third loop were subsequently identified and deleted (atg9 489-502; atg9 E491,4A). The aforementioned mutants were tagged at the N-terminus with GFP. We have shown previously that Atg9 is essential for glucose-induced micropexophagy specifically SM expansion. We first screened these mutants for functionality: whether they were able to rescue the R19 insertion mutant (Chang et al. 2005; Schroder and Dunn, 2006). Within 6 h of glucose adaptation, more than 80% of AOX was degraded by wild-type GS115 cells (Figure 5-3). In comparison, only about 40% of AOX was degraded in R19 cells. R19 cells expressing GFP-Atg9 was able to efficiently degrade AOX. A moderate rescue of R19 was observed when expressing atg9 N and atg9 QQHKA with about 50% AOX degraded. atg9 L3, atg9 489-502, atg9 579-668/ QQHKA and atg9 WEGKY failed to complement R19 as well, with no less than 30% AOX activity remaining. R19 cells expressing atg9 579-668 117

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exhibited less AOX degradation than R19 itself. Interestingly, cells expressing the atg9 579668/ QQHKA double mutant degraded AOX bette r than cells expressing atg9 579-668 alone. None of the atg9 mutants supported AOX degradati on as seen in wild-type cells, implying that all the domains were essential for Atg9 function in glucose-induced micropexophagy. We had also shown that Atg9 is required for nitrogen starvati on-induced proteolysis (Chang et al. 2005). Therefore, we then examined th e ability of cells expressing these atg9 mutants to degrade endogenous protein during nitrogen starvation (Figure 5-4). The atg9 mutants were expressed in cells lacking func tional Atg9 and harboring an arginine auxotrophy (WDY55) to permit amino acid st arvation-induced prot eolysis. When wild-type cells were starved for nitrogen and amino acids, the cellular protein degrad ation average rate was 0.4% per hour. Protein degradation of atg9 lacking cells on average was 0.05% per hour, an 8-fold lower rate than wild-type. Degradati on in cells harboring full-length GFP-Atg9 was similar to that of the wild-type control. WDY55 cells expressing atg9 harboring mutations encompassing regions of the loop between the third and fourth transmembrane domains, atg9 489-502 and atg9 E491,4A, or atg9 579-668 poorly degraded endogenous pr oteins at 0.15% per hour. When atg9-W607A/Y611A or atg9 N were expressed protein degrada tion was only partially restored to almost 0.2% per hour. The data suggest th at these mutants are not fully functional in autophagy induced by amino acid starvation. Atg9 L3 and Atg9 579-668 Mutants Remain In The ER We have shown previously that the loop #3 between transmembrane regions #3 and #4 was required for the exit of Atg9 out of the ER (Chang et al., 2005; Schroder and Dunn, 2006). Because the minimal sequence necessary for atg9 L3 to be retained in the ER was not known, the homology of P. pastoris, S. cerevisiae, D. melanogaster, C. elegans, and H. sapiens was matched. The domain of the highest homology within atg9 L3 is the di-acidic YxDxE motif 118

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(Schroder and Dunn, 2006). This se quence was first localized to the C-terminus of the VSV-G protein; the YKDAD motif is essential for th e cystic fibrosis transmembrane conductance regulator to exit form the ER. The critical 6 amino acids located in the highest homology region were shown to be essential for CFTR from the ER and this is localized to an accessible loop as in the homologous Atg9 sequence. A similar motif is also present in Kir2.1, a potassium channel membrane protein (Schroder and Dunn, 2006). To determine the significance of the di-acidic motif as compared to the larger atg9 L3 mutant, two mutations were made within the region of highest homology. atg9 489-502 obliterated the seque nce altogether. Atg9 E491,4A changed the acidic glutamates at positions 491 and 494 to non-reactive alanines. These atg9 mutants were not functional for micropexophagy or autophagy (see above). After finding that no variant of the atg9 L3 mutant is able to complement R19 cells by AOX assay, the cellular distribution of thes e atg9 mutants duri ng glucose-induced micropexophagy was examined. DMM1 cells expressing GFP-Ppatg9 L3 was found at the ER surrounding the nucleus (Figure 5-5). In a ddition, the cellular location of GFP-Ppatg9 489-502 and GFP-Ppatg9 E491,4A was restricted to the ER n ear the cell surface and surrounding the nucleus (Figure 5-6). These cells showed the same fluorescence distribution as the R19 cells no matter which Ppatg9 L3 variant was expressed. Next, we examined the cellular location of GFP-atg9 579-668 expressed behind the GAPDH promoter. The GFP-atg9 579-668 was present at the peripheral and nuclear ER (Figure 5-7). Thus in addition to the loop region between the third and fourth transmembrane dom ains, residues 579-668 of Atg9 appear to also be an ER exit motif. To verify that the fluorescing region was indeed the ER which surrounds the nucleus in P. pastoris cells, R19 cells harbouring the ER-localiz ing mutations were fixed with DAPI and 119

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observed by fluorescence microscopy (Figure 5-8). Each R19 strain expressing these GFPtagged atg9 mutants exhibited the same mor phology: the blue DAPI-stained nucleus was surrounded by GFP. The preliminary trafficking of Atg9, a transm embrane protein, was never before defined. Hence, the finding that certain atg9 mutants rema in in the ER is noteworthy. Moreover, there appear to be two distinct ER re tention and/or exit motifs complete ly independent of one another. Atg9 QQHKA Remains At The PC9 The QQHKA sequence at the C-terminus of Atg9 was revealed by a database inquiry to be a putative C-terminal ER exit motif. DMM1 cells expressing BFP-SKL and GFPPpatg9 QQHKA behind the GAPDH promoter were visualized during glucose-induced micropexophagy. GFP-Ppatg9 QQHKA localized to one or more punctate cytosolic structures of uniform size at the cell peri phery (Figure 5-9). The GFP-atg9 QQHKA was not visualized at the PVS or the SM. Since the localization of atg9 QQHKA appeared to be restricted to the PC9, we decided to further characterize this compartment by subc ellular gradient fracti onation (Figure 5-10). Under our gradient conditions, the ER marker pCPY and the mitochondrial marker CoxIV-GFP distributed between fractions 9-11, while the vacuole marker CPY and tER marker Sec13 migrated at fractions 3-5. Full-length GFP-Atg9 displayed a bi modal distribution peaking at fractions 7 and 11 in cells grown in methanol alone. Upon switching to glucose, GFP-Atg9 was peaked at fractions 4-6. During glucose adaptation, atg9 QQHKA localized broadly to fractions 7 through 13 with a peak at fraction 9. At this time, atg9 579-668 visualized by fluorescence to be at the ER, localized with the ER marker (fractions 9-11), but was also found at fractions 3 through 7. Based on this data, we suggest that the PC9 co mpartment distributed around fraction 9 lighter in density than the ER and mitochondria, but heav ier than the tER and 120

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vacuole. Upon glucose adaptation, GFP-Atg9 redi stributed from a bimodal distribution to a lighter fraction, which may be the an alogous to the PVS and/or SM. Atg9-W607A/Y611A Fails To Localized To Th e Sequestering Membranes (SM) On The Vacuole The WEGKY domain past the Atg9 last tr ansmembrane domain present at amino acids 607-611 was revealed by database inquiry to be the motif shared by Pex14 and Pex5. Since this motif appears to be the same motif in Pex5 re quired for the interaction of Pex13 and Pex14, we decided to delete the WEGKY site. R19 cells expressing GFP-atg9-W607A/Y611A were examined by fluorescence microscopy (Figure 5-11). Upon glucose adaptation, the WEGKY mutant localized to the cytosolic PC9, the PVS n ear the base of the SM and occasionally at the PAS near the tips of the vacuolar arms but failed to traffic to the SM. We also compared the molecular size of this mutant with normal GFP-Atg9 and the other mutants. Full-length form of GFP-Atg9 migrated as two bands; one at 130kDa, the predicted size, and a second at a sligh tly larger size of about 140kDa (Figure 5-12). Meanwhile, atg9W607A/Y611A migrated as a si ngle band at 130kDa. The slower migrating band was not observed. Upon comparison, the atg9 QQHKA mutant displayed a doubl et; one at the predicted mass and a second unknown band migra ting slightly larger. Atg9 N Localizes To The Vacuole But Is Not Functional By AOX Assay Previous work has shown that the N-terminus of ScAtg9 is vital for its interaction with ScAtg11 (He et al., 2006; Chang and Huang, 2007). Furtherm ore, we have shown that Atg11 is needed for Atg9 transport to the PVS. The data presented in Figures 5-3 and 5-4 showed that the N mutant was defective in both glucose-induced micropexophagy a nd starvation-induced proteolysis. DMM1 cells expressing GFP-PpAtg9 N were adapted from YNM to YND then examined by fluorescence microscopy. GFP-Atg9 N localized to the vacuole and SM (see 121

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Chapter 3). Hence, an atg9 mutant lacking the N-terminus was able to transit to the SM. However, the presence of this mutant at the SM was not sufficient to support micropexophagy. Sites Of Delay In Micropexophagy Caused By The Atg9 Mutants We had observed that in cells lacking Vac8, a vacuolar me mbrane associated protein, micropexophagy was blocked at completion. That is, the SM completely surrounded the peroxisomes, but the peroxisomes were not degraded. However, when cells expressed vac8 ARM4-6 or vac8 ARM5, SM nucleation from th e vacuole was blocked (Fry et al. 2006). Therefore, we decided to determine the sites of micropexophagy blockage in cells expressing the atg9 mutants. In atg9 null cells, the SM could be visualized around the peroxisomes, but the MIPA was not formed (Chang et al., 2005). No cells expressing the mutant forms of atg9 exhibited blocks at nucleation or completion. Instead, cells harboring atg9 mutations exhibit vacuole and SM morphologies undergoing varyin g degrees of expansion based on the SM extensions around the peroxisomes. In atg9 N mutants, micropexophagy appeared blocked at early expansion (see Chapter 3), with atg9 579-668, atg9 L3, and atg9 QQHKA mutants blocked at early or late expansion (see Figure 5-5, 5-7, and 5-9), and finally atg9(W607A/Y611A) cells showing a block during la te expansion (see Fi gure 5-11). These are differences are based on observations of the extents of sequestration by the vacuole and SM relative to the BFP-tagged peroxisomes. The effects of these mutants on MIPA assembly that occurs during late expansion ha s not yet been examined. Discussion In order to better characterize the molecular role of Atg9 in pexophagy, we have set out to define those amino acid motifs responsible for the trafficking and function of Atg9. We have identified a number of motifs that are required for exiting the ER, the PC9, and the PVS. There exist two domains required for At g9 to exit the ER (aa 489-502 specifically E491 122

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and E494 and aa 579-668). The C-terminal QQHKA is necessary for Atg9 to exit the PC9, while W607 and Y611 are essential for Atg9 to become associated with the SM. None of these mutants is functional during glucose-induced micropexophagy or during starvation-induced proteolysis. Finally, the N-term inus is required for function, but not for trafficking of Atg9 to the SM. Our data demonstrates that the specif ic Atg9 domains are essential for its trafficking and function during micropexophagy. Furthermore, the association of Atg9 with the SM is critical for its function. ER Exiting Motifs Of Atg9 Atg9 L3 remains in the ER. The minimal seque nce required for cy cling of Atg9 from the ER was narrowed down by sequential site-directed mutatgenes is to be the E491 and E494. These glutamates are the most highly conserved residues in this region. The importance of this motif to the early trafficking of Atg9 and its high homology emphasizes that the putative loop between the third and fourth transmembrane dom ains is cytosolic. Th e localization of GFPatg9 QQHKA to the PC9 compartment indicates that th e extreme end of the C-terminus is also cytosolic. The requirement for the N-terminus in interactions with ScAtg11 for ScAtg9 to localize to the PAS and the very ear ly early expansion block for atg9 N mutants supports a cytosolic N-terminus model. Thus, both the pu tative Nand C-terminus appear cytosolic, and the predicted 5 or 6 transmembrane domains le nds toward six transmembrane domainsan even number for both termini to be cytosolic. Howe ver, the ER exit motif just past the last transmembrane domain and the presence of the WX XXF/Y motif in the same area, both in the vicinity of the least predicted sixth transmembrane domain, argue against the existence of this sixth transmembrane region, unless the re gion is present somewhere else. Our data also show that Atg9 579-668 fails to exit from the ER. Multiple regions of interest were found upon inspection of the deleted sequence, which is C-terminal of the last 123

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putative transmembrane domain. The WEGKY pentapeptide motif itself was not considered to be the critical motif as point mutations within this sequence led to a phenotype other than ER localization. Within this region there exists a DxE export signal (a.a. 588-592, D P E V S a.a. 606608 E W E a.a. 558-562 D FFR E F S ) (Barlowe, 2003) is a di-acidi c motif involved in recognition by Sec24 for incorporation into the COPII complex via Sec2 3-Sec24 complex interaction (Votsmeier and Gallwitz, 2001; Barlowe, 2003). The dihydrophobic FF/YY/LL/FY motif (a.a. 559-563 FF RE F ) occurs in the cytoplasmic domains of the transmembrane cargo p24 family members as well as the ERGIC53/Emp47 and th e Erv41/46 complexes and is thought to be required for their transport from the ER and th roughout the cell (Sato, 2004) Interestingly, the most highly conserved region when comparing P. pastoris and S. cerevisiae are amino acids 600624 which ends with a tyrosine at amino acid 624 and contains 2 other tyrosines about 5 amino acids apart in both directions. A series of these tyrosines composes the motif localized to the tail of the ERGIC53/Emp47 family and is thought to be important for these transmembrane proteins transport from the ER (Sato, 2004). Thus, future studies are necessary to better define this domain. These ER-exit motifs would likely function at the cytoplasmic surface of the ER possibly interacting with newly forming CO PII vesicles. Evidence suggests that Sec24 has a major role in the recognition of cargo to be packaged into these vesicles. However, Sec23 and Sar1 have also been implicated in cargo recognition and transp ort of proteins out the ER (Sato and Nakano, 2007). Further studies are needed to identify those proteins that interact with these domains thereby promoting Atg9 exit from the ER. PC9 Exiting Motif Of Atg9 The QQHKA domain present at the C-terminal tail of Atg9 is homologous to a KKXXtype ER retention motif. The KKXX C-terminal mo tif retains transmembrane proteins in the ER 124

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and directs their retrograde trafficking from the cis -Golgi in yeast (Gaynor et al. 1994). Disrupted KKXX motifs are transported to the vacuole and proteolytic ally cleaved by Pep4 (Gaynor et al. 1994). In fact, when only one lysine is mutated to glutamine in Wbp1 (a protein required for N-linked glycosylati on of proteins in the ER) it is transported to the vacuole (Gaynor et al. 1994). However, atg9 QQHKA was able to exit the ER to reside at the PC9. During glucose-induced micropexophagy, atg9 QQHKA failed to exit the PC9 compartment. Thus, the dilysine requirement may be very rigid for this motif; a second a lternative is that the QQHKA domain may not be sufficient alone for ER retention. Self-oligomerization enables proteins containing the KKXX domain such as p24 to exit the ER (Sato et al. 2003). Differential binding mechanisms discriminate between ER-resident tr ansmembrane proteins harbouring the KKXX domain and p24 proteins. Thus, even if Atg9 harbours a putative ER retrieval motif, its QQHKA domain could be supers eded if Atg9 is present in a dimeric form. The QQHKA domain is required for Atg9 to exit the PC9. Atg9 may be activated by or interact with another protein vi a this motif. We have shown that Vps15 and Atg11 are required for Atg9 to transit from the PC9 (Chang et al., 2005). Vps15 is a subunit of the PI3K complex and appears to mediate SM nucleation and e xpansion in pexophagy. Atg11 mutants exhibit arrest during during SM expansion (Chang et al., 2005). Atg11 interacts w ith Atg9, but at its Nterminus. No direct interaction has been s hown to exist between Atg9 and Vps15. However, two proteins may act downstream of Vps15 to interact with the QQHKA domain of Atg9, thus explaining the requirement for Vps15 in early Atg9 trafficking. The se rine/threonine kinase Vps15 activates the PI-3 kinase Vps34 which regulates PtdIns(3)P levels. The PtdIns(3)P binding proteins Atg18, Atg20, Atg21, Atg24 are known to be required for autophagy. Atg18 is required for a nucleation event and has no affect on Atg9 trafficking (Stromhaug et al. 2001; 125

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Chang et al., 2005). Atg21 is required for the lipidati on of Atg8 and thus the formation of the MIPA, a late expansion event (Stromhaug et al. 2004). Atg24 is required for completion (Ano et al. 2005). Little is known a bout Atg20 except that it forms a complex with Atg11 and Atg24 at the PAS. Atg20 contains a PX domain, a doma in involved in protein sorting and membrane trafficking that interacts with PI3-P (Yorimitsu and Klionsky, 2005). Atg20 seems to be a good candidate responsible for the tr ansmembrane protein Atg9s trafficking from the PC9. Further studies are needed to evaluate these possibilities. PVS Exiting Motif of Atg9 The WEGKY motif is a WXXXY/F domain within Atg9. Starvation-induced macroautophagy in addition to glucose-induc ed micropexophagy have been shown to be defective in atg9-W607A/Y611A. atg9-W607A/Y611A localizes to the PC9, the PVS and the PAS but fails to associate with the SM dur ing glucose-induced micropexophagy. Proteins sequester at the PVS prior to th eir assembly into the expanding SM and the PAS is thought to be an organization site for the expansion of the vacuolar membranes during MIPA formation (Dunn et al. 2005). The peroxisome membrane protein, Pex14, contains the WXXXY/F domain interacts with the seven WXXXY/F domains of Pe x5 (peroxisome import receptor) (Saidowsky et al., 2001). The N-terminus of Pex14 has been shown to be required for macropexophagy in H. polymorpha. Pex14 has also been shown to be the so le component of the peroxisomal translocon required for pexophagy (Zutphen et al. 2007). The N-terminus of Pex14, a coiled-coil membrane protein, could bind to the WEGKY domain of Atg9. This action might be stabilized by interactions between the co iled-coil regions of Pex14 and Atg11. A second peroxisome membrane protein, Pex13, also interacts with the WXXXF/Y motifs of Pex5 (Otera et al. 2002). However, Pex13 has been shown not to be required for pexophagy (Zutphen et al., 2007). 126

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Nevertheless, it is possible that Atg9 and Atg11 may interact with the Pex13/Pex14 complex thereby directing the movements of the SM around the peroxisomes. We have observed that during glucose-induc ed micropexophagy GFPAtg9 migrates as a doublet on Western blots. However, a doublet is not observed with atg9-W607A/Y611A. The slower migrating band would be consistent with a tyrosine phosphorylation within the WEGKY motif. Two likely protein tyrosine kinases ar e Yak1 and Kns1. Yak1 is involved in glucose signaling and Kns1 has been shown to interact with Atg1 (Hartley et al. 1994; Ptacek et al. 2005; Pratt et al. 2007). Further studies are necessary to determine whether either of these proteins phosphorylate Atg9. Functional Domain Of Atg9 During micropexophagy, Atg9 N properly trafficks to the sequestering membranes of the vacuole but pexophagy does not proceed. Thus, th e deletion of the N-terminus is presumably required for Atg9 function in a la te stage of micropexophagy. Thus, the interaction of Atg9 with another protein as mediated by the N-terminus would occur late in micropexophagy. The Nterminus of ScAtg9 has been shown to bind to the second and the first coiled-coil regions of ScAtg11 (He et al., 2006; Chang and Huang, 2007). The PAS is essential for MIPA formation (Dunn et al., 2005). Thus, the interaction of Atg11 with Atg9 may be critical for a late micropexophagic step. Atg24 may also require Atg9 function for membrane fusion events during micropexophagy. Pexophagosomes do not fuse with the vacuole during micropexophagy in atg24 cells (Ano et al., 2005). An interaction between Atg9 and Atg24 mediated by Atg9s Nterminus could explain this defectthe action of the two proteins together may be required for the final step of vacuolar engulfment. 127

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Summary The appearance of the SM in cells harboring mutant Atg9 emphasizes that Atg9 is required for early and late e xpansion during micropexophagy. Furthermore, cells expressing mutant forms of Atg9 were defective in expa nsion, not nucleation or completion. Trafficking mutants of Atg9 did not support micropexophagy. Thes e mutants failed to associate with the SM suggesting that Atg9 functions at th e SM. We have also shown that there exists a domain within the N-terminus that is required for Atg9 functi on at the SM. The data demonstrate that the movements of Atg9 to the SM are essential for SM expansion. 128

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Table 5-1: Pichia pastoris strains Name Genotype Reference GS115 his4 (Cregg et al. 1985) PPF1 (PPY12) arg4 his4 (Yuan et al. 1997) SMD1163 his4 pep4 prb1 (Tuttle and Dunn, 1995) DMM1 GS115::pDM1 (PAOX1BFP-SKL, ZeocinR) (Kim et al. 2001) R19 GS115 atg9-1:: Zeocin (Stromhaug et al., 2001) LAM2 PPF1 atg9::ARG4 This study WDY55 arg4 his4 atg9-1::ZeocinR This study WDY100 PPF1 his4 arg4::pWD26 (PGAPDH RFP-ATG17, ARG4) This study TC10 R19 his4 ::pTC1 (PGAP GFP-ATG9, HIS4) (Chang et al. 2005) AJM22 GS115 his4 ::pPS69(PGAPDH GFP/HA-ATG2, HIS4) This study AJM25 AJM22::pAJM6 (PGAPDH mRFP-ATG9, ZeocinR) This study AJM26 GS115 his4 ::pAJM6 (PGAPDH mRFP-ATG9, Zeocin) (Chang et al. 2005) AJM32 AJM26 his4 :: pPS64 (PGAPDH GFP/HA-ATG11, HIS4) (Chang et al. 2005) AJM33 AJM26 his4 :: pPS55-G12 (PGAPDH GFP/HA-ATG18, HIS4) This study ASK4 R19 his4 ::pGFP-G14( L3) (PGAP GFP-atg9 L3, HIS4) This study ASK5 R19 his4 ::pGFP-G14( N) (PGAP GFP-atg9 N, HIS4) This study ASK6 R19 his4 ::pGFP-G14( C) (PGAP GFP-atg9 C, HIS4) This study ASK7 DMM1 his4 ::pGFP-G14( L3) (PGAP GFP-atg9L3, HIS4) This study ASK8 DMM1 his4 ::pGFP-G14( N) (PGAP GFP-atg9N, HIS4) This study ASK9 DMM1 his4 ::pGFP-G14( C) (PGAP GFP-atg9C, HIS4) This study LAM6 R19 his4 ::(PGAPDH GFP-atg9 QQHKA, HIS4) This study LAM7 R19 his4 ::(PGAPDH GFP-atg9 579-668, HIS4) This study LAM8 R19 his4 ::(PGAPDH GFP-atg9 579-668 / QQHKA, HIS4) This study LAM9 DMM1 his4 ::(PGAPDH GFP-atg9 QQHKA, HIS4) This study LAM10 DMM1 his4 ::(PGAPDH GFP-atg9 579-668, HIS4) This study LAM11 DMM1 his4 ::(PGAPDH GFP-atg9 579-668 / QQHKA, HIS4) This study LAM19 R19 his4 ::(PGAPDH GFP-atg9 489-502, HIS4) This study LAM20 DMM1 his4 ::(PGAPDH GFP-atg9 489-502, HIS4) This study LAM36 R19 his4 ::(PGAPDH GFP-atg9 W607A/Y611A, HIS4) This study LAM51 R19 his4 ::(PGAPDH GFP-atg9 E 491,4A, HIS4) This study LAM53 AJM26 his4 ::pWD24 (PGAPDH GFP-ATG17, HIS4) This study LAM61 WDY100 his4 ::(PGAPDH GFP-atg9 QQHKA, HIS4) This study LAM62 WDY100 his4 ::(PGAPDH GFP-atg9 W607A/Y611A, HIS4) This study LAM65 WDY55 his4 arg4 atg9::pTC1 (PGAPDH GFP-ATG9, HIS4) This study WDY75 AJM26 his4 ::pWD21 (PGAPDH GFP-PpATG8, HIS4) This study 129

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Figure 5-1. Localization of Atg9, Atg11 and Atg17 to the PVS. LAM25, LAM32, LAM33, LAM53, and WDY75 were grown in YNM, adapte d to YND for 2 hours, and the Atg proteins 130

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visualize by fluorescence microsc opy. RFP-Atg9p is found at a pe ripheral compartment without Atg2 (white arrowhead), but with Atg17p (yellow arrowhead). RFP-Atg9p is also present at the PVS (white arrows) colocali zed with Atg11p and Atg17p (ye llow arrows). GFP-Atg8p is localized to the PAS and MIPA (green arrows). 131

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Figure 5-2. Linear structure of the PpAtg9 tr ansmembrane protein and deletions performed by site-directed mutagenesis Atg9 is an 885 amino acid long protein required for micropexophagy. The linear representation of Atg9 and its muta ted regions are drawn to scale. GFP is Nterminally attached to Atg9. There exist 5-6 putative transmembrane (T M) domains. Five of putative TM domains are shown: L227-I244 (18 a. a.), L277-L295 (19 a.a.), L439-L463 (25 a.a.), F528-L546 (19 a.a.), and P558-C576 (19 a.a.), and are drawn to scale. Numerous motifs have been identified in situ WLFSF (a.a. 235-239) and WEGK Y (a.a. 607-611) are WXXXF/Y motifs shared with Pex5 and Pex14, although the WL FSF motif may be mask ed within the first transmembrane domain. The last 5 C-terminal amino acids are QQHKA (a.a. 881-885). Sitedirected mutagenesis produced atg9 N, atg9 L3 (the region between the third and fourth transmembrane domain), atg9 489-502 (a highly conserved region of the region mutated in atg9 L3), atg9 579-668 (a conserved region after the last transmembrane domain), atg9W607A/Y611A (the WXXXF/Y motif sh ared by Pex5 and Pex14), and atg9 QQHKA. GFP is attached to the most N-terminal region of each mutant. 132

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Figure 5-3. Atg9 mutants fail to su pport glucose-induced micropexophagy Wild-type, atg9 null mutants, and null mutants expressing mutant forms of atg9 were grown in YNM and adapted to YND. AOX levels were measured at 0 and 6 hours post glucose adaptation. The values were expressed as the percentage of AOX activity remaining (average SEM). 133

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Figure 5-4. Protein degrada tion of Atg9 mutant cells during nitrogen starvation Wild-type, atg9 null mutants, and null mutants expressing mutant forms of atg9 were grown in minimal medium with 14C-valine. After 18 hours, the cells we re switched to nitrogen starvation medium and protein degradation measured as previously described in Materials and Methods. 134

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Figure 5-5. GFP-atg9 L3 remains in the ER during glucose-induced micropexophagy. DMM1 cells expressing BFP-SKL and GFP-atg9 L3 were grown in YNM in the presence of FM4-64. After 18 hours, the cells were adapted to YND for 2 hours and visualized by fluorescence microscopy. The GFP-atg9 L3 did not colocalize with th e FM4-64 labeled vacuole or sequestering membranes that surrounded the BFP labeled peroxisomes. Instead, the GFPatg9 L3 appeared at the cell periphery and around the nucleus (arrows). 135

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Figure 5-6. GFP-atg9 489-502 and GFP-atg9 E491,4A remain in the ER during glucoseinduced micropexophagy. R19 cells expressing GFP-atg9 489-502 or GFP-atg9 E491,4A were grown in YNM in the presence of FM4-64. Afte r 18 hours, the cells were adapted to YND for 2 hours and visualized by fluoresce nce microscopy. The GFP-atg9 489-502 or GFPatg9 E491,4A did not colocalize with the FM4-64 labe led vacuole or sequestering membranes. Instead, the GFP-tagged proteins appeared at the cell periphery and around the nucleus. 136

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Figure 5-7. GFP-atg9 579-668 remains in the ER during glucose-induced micropexophagy. DMM1 cells expressing BFP-SKL and GFP-atg9 579-668 were grown in YNM in the presence of FM4-64. After 18 hours, the cells were adapted to YND for 2 hours and visualized by fluorescence microscopy. The GFP-atg9 579-668 did not colocalize with the FM4-64 labeled vacuole or sequestering membrane s that surrounded the BFP labele d peroxisomes. Instead, the GFP-atg9 579-668 appeared at the nuclear membranes (arrows). 137

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Figure 5-8. GFP-atg9 489-502, GFP-atg9 E491,4A, and GFP-atg9 579-668 localized to the nuclear membranes. To confirm that GFP-atg9 489-502, atg9 E491,4A, and atg9 579-668 all localize to the P. pastoris ER which surrounds the nucleus a nd not to another organelle, the nuclear DNA was stained with 4',6-diamidino-2phenylindole (DAPI). R19 cells expressing GFP-atg9 489-502 or GFP-atg9 E491,4A or GFP-atg9 579-668 were grown in YNM in the presence. After 18 hours, the cells were ad apted to YND for 2 hours and formaldehyde fixed. After staining the cells with DAPI, they were visualized by fluorescence microscopy. The GFPtagged proteins were positioned at the nuclear membrane ar ound the DAPI stained DNA. 138

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Figure 5-9. GFP-atg9 QQHKA remains at peripheral vesi cles during glucose-induced micropexophagy. DMM1 cells expressing BFP-SKL and GFP-atg9 QQHKA were grown in YNM in the presence of FM4-64. After 18 hours, the cells were adapted to YND for 2 hours and visualized by fluorescence microscopy. The GFP-atg9 QQHKA did not colocalize with the FM4-64 labeled vacuole or sequestering me mbranes that surrounded the BFP labeled peroxisomes. Instead, the GFP-atg9 QQHKA localized to peripheral vesicles (arrows). 139

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Figure 5-10. Distribution of Atg9 vesicles on sucrose gradients. TC10 were either grown in YNM for 18 hours (A) or adapted from YNM to YND for 4 hours (B). LAM7 was adapted from YNM to YND for 4 hours (C) and LAM6 was grown in YNM for 18 hours. The cells were spheroblasted, lysed, and the organelles sedime nted at 100,000 xg. The pellet was resuspended and loaded onto a continuous sucrose gradient (15 55%). The gradient was centrifuged to equilibrium at 25k rpm for 15 hours. The ER marker pCPY and the mitochondrial marker CoxIV-GFP distribute around fractions 9-11. CPY which localizes to the vacuole and Sec13 which localizes to the transition endoplasmic reticulum (tER) di stributed around fractions 4 and 5. 140

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Figure 5-11. GFP-atg9(W607A/Y6 11A) fails to localize to the sequestering membranes (SM) during glucose-induced micropexophagy. R19 cells expressing GFP-at g9(W607A/Y611A) were grown in YNM in the presence of FM4-64. Afte r 18 hours, the cells were adapted to YND for 2 hours and visualized by fluorescence microscopy. GFP-atg9(W607A/Y611A ) localized to the PC9 (yellow arrowhead), PVS (blue arrow) and PAS (white arrow), but not the sequestering membranes or vacuole. 141

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142 Figure 5-12. GFP-Atg9, not GFPatg9(W607A/Y611A), migrates as a doublet on Westerns. At 3, 6, 9 and 12 hour post glucose adapta tion, cells expressing GFP-Atg9, GFPatg9(W607A/Y611A), or GFP-atg9 QQHKA, were lysed and the molecular sizes of the GFPtagged proteins evaluated on Western blots. Both GFP-Atg9 and GFP-atg9 QQHKA migrated as a doublet: one band at 130kDa (arrow), the pred icted size, and a second band at a slightly larger size of about 140kDa (asterisk). Meanwh ile, atg9-W607A/Y611A migrated as a single band at 130kDa (arrow).

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CHAPTER 6 SAR1P AND PEXOPHAGY Introduction Autophagy is a lysosomal degradation path way meaning self-eating. Autophagy can be subclassified based on substrate selectivit y and mode of sequestra tion (for reviews see (Levine and Klionsky, 2004; Dunn et al., 2005; Yorimitsu and Klionsky, 2005)). Nonselective microand macroautophagy can be stimulated by nitrogen and amino acid starvation. During microautophagy, sequestration occurs by invagi nations of the lysosomal membrane. Sequestration of macromolecules and orga nelles into autophagosomes proceeds by macroautophagy. These autophagosomes eventually fuse with the lysosomes thereby delivering their cargo for degradation. The se lective degradation of peroxisomes has been referred to as pexophagy. Micropexophagy is the seque stration of peroxisomes by th e vacuole for degradation. When Pichia pastoris are subjected to glucose adapta tion, clusters of peroxisomes are surrounded by finger-like prot rusions originating from the vacuole. The peroxisome cluster is then incorporated into the vacuolar lumen where it is subsequently degraded. During adaptation from methanol to ethanol, individual peroxi somes are sequestered in to autophagosomes and delivered to the vacuole for degradat ion. This process is called macropexophagy. During glucose-induced micropexophagy, nucle ation of the sequestering membranes proceeds from the vacuole. Membrane nucleation requires Atg18p6, Vac8p and Vps15p(Stasyk et al. 1999; Guan et al., 2001; Fry et al. 2006). Afterwards, the sequestering membranes expand to surround the peroxisomes. This expa nsion includes membranes from the vacuole and other undefined membranes. Membrane expansion requires Atg2p, Atg9p and Atg11p (Kim et 6 We will conform to the ATG nomenclature. Unless stated by designations for other species, we refer to the P. pastoris homologs of the Sar1, Atg and Sec proteins in this paper. 143

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al. 2001; Stromhaug et al., 2001; Chang et al., 2005). We have demonstrated that Atg9p, an integral membrane protein, traffics from a unique, peripheral compartment (PC9) 7 to perivacuolar structures (PVS) a butting the vacuole, and finally to the sequestering membranes. The movement of Atg9p to the sequestering membranes requires Atg2p and Atg11p. Mukaiyama, et al. have characterized th e micropexophagic membrane apparatus (MIPA) required for expansion and completion events of micropexophagy (Mukaiyama et al., 2004). The de novo assembly of the MIPA is positioned be tween expanding sequestering membranes. The assembly of the MIPA from a pre-autoph agosome structure (PAS) requires Atg7p, Atg8p, and Atg9p (Mukaiyama et al., 2004; Chang et al., 2005). Atg8p, Atg9p and Atg17p localize to the PAS, with Atg8p also being found at the MIPA (Mukaiyama et al., 2004; Chang et al. 2005). Finally, completion of micropexophagy pro ceeds upon the fusion of MIPA with the adjacent sequestering membranes and incorporati on of the peroxisomes into the vacuole for degradation. These fusion events require Vac8p and Atg24p (Ano et al., 2005; Fry et al. 2006; Oku et al., 2006). During ethanol-induced macropexophagy, peroxi somes are enwrapped by two or more membranes of unknown origin forming pexophagosomes (Dunn et al., 2005). The molecular events responsible for the formation of th e pexophagosome remain rather vague. These sequestering membranes contain Atg8p and Atg25p which arise fr om the PAS that contain a number of Atg proteins, in cluding Atg8p, Atg9p and Atg17p (Chang et al. 2005; Oku et al., 2006). The completion of the pexophagosome requires Atg8p and Atg21p. The outer 7 The abbreviations used are: AOX, alcohol oxidase; COPII, coat protein complex II; ER, endoplasmic reticulum; FM4-64, N-(triethlyammoniumpropyl)-4-(p-diethylaminophenylhexatrienyl) pyridinium dibromide; MIPA, micropexophagy-specific membrane apparatus; PAS, pr e-autophagosome structure; PC2, Atg2, peripheral compartment; PC9, Atg9 peripheral compartment; PI4P, phosphatidylinositol 4-monophosphate; PtdEtn, phosphatidylethanolamine; PVS, perivacuolar structure; SM, sequestering membranes; TCA, trichloroacetic acid; and v-SNARE, vesicle solubl e NSF attachment receptor. 144

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membrane of the pexophagosome then fuses with the vacuole delivering the peroxisomes for degradation. In addition to Atg25p at the pexophagosome, fusion requires Vam7p present at the vacuole and Atg24p present at th e vertices and boundary regions of the fusion complex (Ano et al. 2005; Monastyrska et al. 2005; Stevens et al. 2005). Within the vacuole, the inner membrane of the pexophagosome is ruptured by the lipase Atg15p and the released peroxisomes are rapidly degraded by proteinases (Teter et al. 2001). Recent data suggest that a number of molecula r events of the secretory pathway may be shared by autophagy. For example, ScAtg8p in teracts with the v-SNARE proteins Bet1p, Sec22p, and Nyv1p, which are required for endoplasmic reticulum (ER) to Golgi trafficking and vacuolar protein traffi cking (Legesse-Miller et al. 2000). In addition, studies have shown that a number of Sec proteins are required for autophagy, namely ScSec16p, ScSec17p, ScSec23p, ScSec24p, ScSec31p, ScSec12p, ScSec18p, and ScSec7p (Ishihara et al. 2001; Hamasaki et al. 2003; Reggiori et al. 2004). The roles for ScSec16p, ScSec17p and ScSec23p have yet to be clearly defined (Ishihara et al. 2001). ScSec12p facilitates the formation of the autophagic vacuole from the PAS (Reggiori et al. 2004). When sec24ts and sec31ts mutants are exposed to nonpermissive temperatures, ScAtg8p is found at cy tosolic structures, which are either multiple PAS forming autophagic vacuoles or autophagic vacuoles failing to fuse with the vacuole. Finally, ScSec18p and ScSec7p appear to be requ ired for the fusion of the autophagic vacuole with the vacuole (Ishihara et al., 2001; Reggiori et al. 2004). Hence, a number of Sec proteins are indispensable for both prot ein secretion and autophagy. Some of the secretory proteins required for autophagosome formation have a role in COPII-mediated trafficking (Ishihara et al., 2001; Hamasaki et al. 2003; Reggiori et al. 2004). ScSec12p is a GDP/GTP transmembrane exchange factor which influences ScSar1p activity (for 145

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review see (Sato and Nakano, 2007). ScSar1p, st imulated by the exchange of GTP for GDP, binds to the ER membrane. ScSec23p inter acts with ScSec24p, whic h bind to ScSar1p. ScSec13p will bind to the ScSar1p-ScSec23p/ ScSec24p coat protein complex promoting vesicular budding. Point mutations in ScSar1p inhibit COPII-dependent protein sorting (Ward et al. 2001). The ScSar1pT39N mutant has a high affinity for GDP and blocks COPII vesicle formation from the ER. The ScSar1pH79G mutant is unable to hydrolyze GTP and prevents disassembly of COPII vesicle coats thereby inhib iting retrieval of proteins to the ER. The data suggest that COPII-dependent protein traffi cking is essential for ongoing autophagy and may implicate the ER in the assembly of the autophagosomes (Ishihara et al. 2001; Hamasaki et al. 2003; Reggiori et al. 2004). Studies in S. cerevisiae have shown that the COPII comp lex of proteins is required for starvation-induced autophagy. However, the role of this protein complex in pexophagy has not been evaluated. In this study, we characterized the essential role of Sar1p in microand macropexophagy in Pichia pastoris We demonstrated that when dominant-negative mutants, Sar1pT34N or Sar1pH79G, are expressed both microand macropexophagy are repressed. The nucleation and amplification of the sequesteri ng membranes appeared to proceed normally during micropexophagy. However, the assembly of the MIPA was impaired when these Sar1p mutants were expressed. The failure of MIPA to form was not due to in correct sorting of Atg2p, Atg9p, or Atg11p or an inactivation of Atg7p. Howe ver, the lipidation of Atg8p appeared to be altered. During micropexophagy, the pexophagosome failed to assemble when Sar1pT34N was expressed. To the contrary, the expression of Sar1pH79G did not appear to alter the formation of the pexophagosome, but disrupted its delivery of the peroxisomes to the vacuole. Our data 146

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suggest that Sar1p is essential fo r the trafficking of ER proteins into and out of the sequestering membranes of microan d macropexophagy. Materials and Methods Yeast Strains And Media The yeast strains used in this study are listed in Table 1 and were routinely cultured at 30 C in YPD (1% Bacto yeast extract, 2% Bacto peptone, and 2% dextrose). P. pastoris was grown in YNM (0.67% yeast nitrogen base, 0.4 mg/l biotin, and 0.5% methanol) to induce peroxisome biogenesis. The degradation of peroxisomes was induced when cells were transferred from YNM to YND (0.67% yeast nitr ogen base, 0.4 mg/l biotin, and 2% glucose) or YNE (0.67% yeast nitrogen base, 0.4 mg/l bio tin, and 0.5% ethanol). Nitrogen starvation medium, SD(-N), contained 0.17% yeast nitrogen base (without amino acids and NH4SO4) and 2% glucose. All media contained 2% agar when made as plates. His tidine and arginine were added at 40 g/ml when needed. Vector amplification was done in E. coli (DH5 ) cultured at 37 C in LB (0.5% Bacto yeast extract, 1% Bact o tryptone, and 1% NaCl) with ampicillin (100 g/ml). Zeocin was added at 25 g/ml when culturing DH5 and 100 g/ml when culturing P. pastoris. Quantitative Assessment Of Alcohol Oxidase (AOX) Degradation Cells were grown in 20 ml of YNM with metha nol as sole carbon and energy source. At 40 hours, 0.4 g glucose was added with or without 0.1 mM copper sulfat e. Aliquots (2 ml) of cells (8.0 OD600) at 0 and 6 hours of glucose adaptation we re pelleted and resuspended in 1 ml of 20 mM Tris pH 7.5 containing 50 mM NaCl, 1 mM EDTA, 1 mM PMSF, 1 g/ml pepstatin A, and 0.5 g/ml leupeptin. The cells were then lysed by vortexing in the presence of 0.5 ml glass beads (425-600 microns). The glass beads and cellular debris were removed by centrifugation 147

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and AOX was measured by adding 50 l of this extract to 3 ml of reaction mix containing 3.4 U/ml horseradish peroxidase and 0.53 mg/ml 2,2-azino-bis(3-ethylbenz-thazoline-6-sulfonic acid) in 33 mM potassium phosphate buffer pH 7.5. The reaction was initiated by adding 10 l methanol and terminated after 20 minutes by adding 200 l of 4N HCl and the absorbance read at 410 nm. Measurements of Protein Degradation The degradation of cellular proteins during nitr ogen starvation was performed as described previously (Tuttle and Dunn, 1995). Endogenous proteins were labeled with 1 Ci/ml 14C-valine for 16 hours in 0.67% yeast nitrogen base 2% glucose, 0.4 mg/l biotin, and 40 g/ml histidine (if necessary). The cells were then washed and switched to SD(-N) with or without 0.1 mM copper sulfate and supplemented with 10 mM valine. Aliquots were removed at 2 24 hours of chase and trichloroacetic acid (TCA) was added to a fi nal concentration of 20%. Acid soluble and insoluble radioactivity was sepa rated by centrifugation and quantif ied by scintillation counting. The rates of protein degradation were calculated from the slopes of the linear plots of TCAsoluble radioactivity ve rsus time of chase. Western Blot Analysis Cells were prepared for SDS-PAGE and West ern blots as previous ly described (Chang et al. 2005). The cells (2-3 OD600) were pelleted, washed with water, and lysed in 150 l of SDS sample buffer (67 mM Tris, pH 6.8, 2% SDS, 10% glycerol, 0.1% bromophenol blue, 1.5% DTT solution containing 4 mM PMSF, 4 mM pepstati n A, and 2 mM leupeptin) by vortexing with glass beads (425-600 micron diameter). Just prio r to SDS-PAGE, the samp les were incubated at 100 C for 5 minutes and cell debris and glass be ads were removed by centrifugation. Five to twenty microliters of sample were loaded onto 8%, 10% or 15% polya crylamide gels. Upon 148

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electrophoresis, the proteins were transferred to nitrocellulose using the Trans-Blot SD-Dry Transfer Cell (Bio-Rad Laboratories, Hercules, CA ). The blots were blocked in 5% nonfat dried milk in PBS containing 0.1% Tween 20 (PBST) a nd incubated with the pr imary antibodies rabbit anti-AOX (Tuttle and Dunn, 1995), rabbit antiHA (Covance, Emeryville, CA) or rabbit antiGFP (Sigma, St. Louis, MO). The blots were then washed in PBST, and followed by incubation with secondary antibodies conjuga ted to HRP. After washing w ith PBST, the antibodies were detected using ECL (Amersham, Piscataway, NJ). Construction of Conditional Expression Vectors Expression vectors containing sar1 (T34N) and sar1(H79G) behind the copper-inducible CUP promoter were constructed as follows. PpARG4 gene was amplified by PCR from pYM30 (kind gift of Dr Jim Creg g, Keck Graduate Institute) and insert ed into the SspI and NdeI sites of pUC19. The resulting plasmid, pUC19-PpArg4, wa s amplified, cut with NdeI, blunt-ended with T4 DNA polymerase, and digested with EcoRI. The S. cerevisiae CUP promoter was excised from pCAJ3 vector with BamHI (blunt-ended w ith T4 DNA polymerase) and EcoRI, and then ligated into the pUC19-PpArg4. Next, the poly-A sequence from pIB2 was excised and inserted into the HindIII and AflII site yielding the pWD16 conditional expres sion vector. Finally, sar1(T34N) and sar1(H79G) kindly provided by Dr Ben Glic k (University of Chicago) were amplified by PCR and inserted into the EcoRI and HindIII sites of pWD16. GFPATG17 was constructed by inserting ATG17 amplified by PCR from genomic DNA into the KpnI and XhoI sites of pPS55 following similar procedures used to construct expressi on vectors containing GFPATG2 GFPATG8 GFPATG9 GFPATG11 and GFPATG18 (Guan et al., 2001; Kim et al. 2001; Stromhaug et al., 2001; Chang et al., 2005). RFP-HDEL was provided by Dr Ben Glick (University of Chicago) and pAJM6 (PGAPDH mRFP-PpAtg9, ZeocinR) and pWD18 (PATG7 149

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atg7-HA (C518S), HIS4) were constructed as previously described (Yuan et al. 1999; Chang et al. 2005). Yeast Transformation Cells grown overnight in YPD to a density of A600 = 1.0 were harvested and treated with 10 mM DTT in YPD containing 25 mM HE PES, pH 8, for 15 minutes at 30 C. The cells were washed twice in ice-cold water and once in 1 M sorbitol and then resuspended in 1 M sorbitol. Cells (40 l) were mixed with 0.2-1 g of linearized vector and transferred to a 0.2 cm gap cuvette (Bio-Rad, Hercules, CA), and the DNA introduced by electroporation at 1.5 kV, 25 F, 400 (Gene Pulser, Bio-Rad Corp.). The cells transformed with vectors containing the HIS4 gene were transferred to plat es containing 0.67% yeast nitrogen base without amino acids, 2% glucose, 1 M sorbitol, 0.4 mg/l biotin, and 2% agar and incubated at 30 C for 3 5 days before colonies appeared. For antibio tic selection, Zeocin was added to a final concentration of 100 g/ml. Fluorescence Microscopy Cells expressing GFP and/or mRFP fusion pr oteins were grown in YNM for 20 hours. FM4-64 (Molecular Probes, Eugene, OR) was adde d to a final concentration of 20 ug/ml and cells incubated for 12 16 hours. Cells were th en transferred to YND or YNE with or without 0.1 mM copper sulfate. After 2 4 hours, the ce lls were examined live using a Zeiss Axiophot fluorescence microscope. Image capture wa s done using a SPOT camera (Diagnostics Instruments, Inc., Sterling Heights, MI) interfaced with IP Lab software. Electron Microscopy Upon glucose or ethanol adapta tion, cells were harvested by centrifugation, washed in water, and fixed with 1.5% KMnO4 in veronal-acetate buffer ( 30 mM sodium acetate, 30 mM 150

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sodium barbital, pH 7.6) for 20 minut es at room temperature (Veenhium et al ., 1983). The samples were dehydrated by washing with increasi ng concentrations of et hanol followed by two washes with 100% propylene oxide. The cells were then infiltrated with a 50:50 mix of propylene oxide and POLY/BED 812 (Polyscien ces, Inc., Warrington, PA) for 16 hours at 4 C and another 24 hours at 22 C under vacuum. Afterwards, the ce lls were infiltrated with 100% POLY/BED with accelerator 2,4,6-Tri(dimethylaminomethyl)phenol (DMP-30, Polysciences, Inc.) for another 2 days at 22 C under vacuum. The cells were pelleted and incubated at 60 C for 2 days to complete the polymerization of th e resin. The cell pellets were mounted on blocks, sectioned, and prepared for examination on a JE OL 100CX transmission electron microscope. Results Effects Of Dominant-Negative Mutants Of Sar1p On Autophagy And Pexophagy ScSec12p, the guanine nucleotide exchange f actor for the G-protein ScSar1p, has been shown to be required for autophagy in S. cerevisiae (Ishihara et al. 2001; Hamasaki et al. 2003; Reggiori et al. 2004). Since Sar1p is essential for cell growth, the role of Sar1p in autophagy and pexophagy has been difficult to evaluate. To circumvent this technical problem, we have constructed dominant-negative forms of Sar 1p whose expression is regulated by a copperinducible CUP1 promoter of S. cerevisiae Sar1pT34N has a high affinity for GDP and blocks COPII vesicle formation. Sar1pH79G is defec tive in hydrolyzing GTP. In the absence of copper, the culture growth of WDY64 and WDY65 in minimal medium appeared normal (Fig. 610). However, when these cells were exposed to 0.1 mM copper sulfate to promote the synthesis of Sar1pT34N and Sar1pH79G, growth was rapidly and dr amatically reduced (Fig. 6-10). The data suggest that we can tightly regulate the expr ession of these proteins with copper additions. 151

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Using the aforementioned cells, we first examin ed whether the expression of these Sar1p mutants affected starvation-i nduced autophagy (Fig.61A). The degradation of endogenous proteins in cells pre-labeled with 14C-valine was measured in nitrogen-starved cells in the absence and presence of copper. Nitrogen starvation induces the nonsel ective delivery of cellular components to the vacuole by microand macroautophagy in yeast. We compared endogenous proteolysis in WT cells to cells expressing the Sar1p mu tant proteins (Fig. 6-1A). We observed a 25% reduction of proteolysis in st arved WT cells exposed to copper. This was compared to the over 60% reduction in starvation -induced proteolysis in cells whose expression of Sar1pT34N or Sar1pH79G had been induced with copper. The data show that Sar1p is required for starvation-induced macroautophagy. Next, we examined the effects of the Sa r1p mutant proteins on glucose-induced micropexophagy (Fig. 6-1B) and ethanol-induced macropexophagy (Fig. 6-1C). Cells were grown in methanol, adapted to gl ucose in the absence and presen ce of copper for 6 hours and the remaining AOX activity quantified. Less than 20% of the AOX remained in the WT cells regardless of the presence of copper. In the absence of copper, 80% of the AOX was degraded by the WDY64 and WDY65 cells. However, when these cells were incuba ted with copper, only 10% of the AOX was lost during gl ucose adaptation. Finally, cel ls were grown in methanol, adapted to ethanol in the absence and presen ce of copper for 0 24 hours and the remaining AOX detected by Western blotting. We observed that in the ab sence of copper AOX protein was almost completely degraded within 9 hours. However, when Sar1pT34N or Sar1pH79G was expressed, AOX protein remained for up to 24 hours. The data reveal that Sar1p is essential for both microand macropexophagy. 152

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Suppression Of Glucose-Induced Micropexoph agy By Dominant-Negative Mutants Of Sar1p Occurs At A Late Sequestration Event We have shown that the expression of Sar 1pT34N or Sar1pH79G dr amatically inhibits the degradation of peroxisomes induced by glucos e adaptation. We next examined whether these proteins inhibited the events of peroxisome seque stration and/or degradatio n. We first examined the morphology of those cells expressing Sar 1pT34N and Sar1pH79G by electron microscopy. WDY64 and WDY65 were adapted from YNM to YND for 4h in the abse nce or presence of copper (Fig. 6-2). Without copper, the endoplasmi c reticulum appeared as discontinuous sheets at the cell periphery and about the nucleus, while the Golgi ap paratus was not easily observed. The morphological changes of these organelles when the Sar1p mutants were expressed were consistent with the known functions of Sar1p. For example, when Sar1pT34N, which inhibits ER budding, was expressed the ER was extended a ppearing as a continuous sheet at the cell periphery and throughout the cell. To the cont rary, when cells expressed Sar1pH79G, which inhibits recycling from the Golg i apparatus back to the ER, the Golgi apparatus appeared as multiple cisternae. During glucose adaptation in cells not expressing the Sar1p mutants, peroxisomes were surrounded by the vacuole or w ithin the vacuole (Fig. 6-2A and 6-2E). In cells expressing Sar1pT34N (Fig. 6-2B-D) or Sar1pH79G (Fig. 6-2F -H), the peroxisome clusters were situated outside the vacuole suggesting that Sar1p was required for delivery of the peroxisomes to the vacuole and not vacuolar degradation. The events of micropexophagy include: sign aling, sequestration, and degradation. The above data suggest that the Sar 1p mutants were suppressing an ev ent prior to degradation most likely sequestration. The engulfmen t of peroxisomes by sequesteri ng membranes (SM) that arise from the vacuole proceeds via SM nucleation, ex pansion, and completion. These events can be visualized by observing the SM with FM4-64 staining incorporating peroxisomes expressing 153

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BFP-SKL (Fig. 6-11). For these studies, we c onstructed cell lines e xpressing BFP-SKL behind the AOX promoter in cells expressing the Sar1p mu tants. At 2h of glucose adaptation, cells were observed at all phases of micropexophagy. During nucleation, the sequestering membranes extend less than halfway around the peroxisomes. About 15% of the cells examined at 2h of glucose adaptation are at this st age. Expansion of the sequest ering membranes can be observed in over half of the cells when the sequest ering membranes extend 50 to 90% around the peroxisomes. The completion events are ch aracterized by peroxisomes being completely surrounded by the sequestering membranes, but the membranes appear segmented and not completely fused. Over 30% of the cells examined were at this stage. Finally, the degradative stage was detected in 5% of the cells. This stage was characterized by a continuous ring of FM464 labeled membranes surrounding peroxisomes that were no longer clearly delineated due to degradation. The data suggest that the expansion and comple tion events are rate-limiting in micropexophagy. We observed 52% of the cells expressing Sar1pT34N at the expansion stage (Fig. 6-2C), 15% at completion (Fig. 6-2D), and 33% at degradation. In cells expressing Sar1pH79G, 23% of the cells were observed at nu cleation, 48% at expansion (Fig. 6-2G), 20% at completion (Fig. 6-2H), and 9% at degradati on. These results suggest that Sar1p was not required for nucleation by likely for the expa nsion and/or completi on events and possibly degradation. Because peroxisomes were sometimes observed within the vacuole, it is possible that one or more vacuolar lipase s or proteinases were missorted and not arriving at the vacuole. Sar1p Is Essential For The Formation Of The MIPA Our data suggest that Sar1p is critical fo r the expansion and/or completion phases of peroxisome sequestration during glucoseinduced micropexophagy. Expansion of the sequestering membranes require s Atg2p, Atg9p, and Atg11p and the assembly of the MIPA, which requires Atg8p. Completion requires MIPA and other proteins including Vac8p and 154

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Atg24p. During expansion, Atg9p traffics from a unique peripheral compartment (PC9) to become associated with the sequestering memb ranes. This trafficking requires Atg2p and Atg11p. Also at this time, Atg8p becomes associat ed with the newly assembled MIPA, which is positioned between the expanding sequestering membra nes. Therefore, we decided to examine the effects of these Sar1p mutants on the trafficking of Atg2p, At g9p, Atg11p, and Atg8p during glucose adaptation. Cell lines expressing Sar1pT34N or Sar1pH79G and GFP-Atg2p, GFPAtg9p, GFP-Atg11p or GFP-Atg8p were constructed (see Table 1). The cells were adapted from YNM to YND in the absence and presence of copper sulfate and examined by fluorescence microscopy (Figs. 6-3 and 6-4). We first examined the trafficking of Atg11p. During micropexophagy, Atg11p colocalizes with Atg9p at the perivacuolar st ructure (PVS) and SM and is required for the trafficking of Atg9p to the PVS (Chang et al., 2005). In the absence of copper, Atg11p was found at the PVS, vacuole and SM at 2h of gluc ose adaptation. The distribution of Atg11p was unaltered when either Sar1pT34N or Sar1pH79G we re expressed (Fig. 6-3). Next, we examined the distribution of GFP-Atg2p in cells. Upon glucose adaptation, Atg2p becomes associated with foci known as the Atg2p peripheral compartmen t (PC2). Its role in the expansion of the SM is not defined, but Atg2p is required for the transit of Atg9p to the SM. When LAM44 and LAM45 were adapted from YNM to YND in the absence of copper, Atg2p was found in the cytosol and at foci situated at the cell periphery and n ear the vacuole. When cells were adapted to glucose in the presence of copper, Atg2p was virtually abse nt from the cytosol and was instead observed in foci and larger irregularly shaped structures (F ig. 6-3, yellow arrows). These coalescing structures may represent an expansi on of this peripheral compartment due to the 155

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expression of the Sar1p mutants. The data sugg est that the trafficking of Atg2p and Atg11p were not significantly influenced by Sar1p. We have previously shown that during the membrane expansion events Atg9p transits from PC9, to the PVS, and finally to the SM (Chang et al., 2005). Atg9p can also be observed occasionally colocalizing with Atg8p and Atg17p at th e PAS. We next examined the effects of Sar1pT34N or Sar1pH79G on the cellular distri bution of GFP-Atg9p duri ng glucose adaptation. In the absence of copper, GFP-Atg9p was found at the PC9, PVS, and SM (Fig. 6-3). Atg9p was also present at the vacuole membrane. When copper was added, the distribution of GFP-Atg9p was not dramatically altered. In addition to the peripheral foci of the PC9 compartment (Fig. 63, white arrowhead), Atg9p was present at the PVS near the base of the sequestering arms (Fig. 6-3, white arrow), the PAS or MIPA nucleation site (Fig. 6-3, yellow arrowhead), and the SM. We have reported that Atg2p and Atg9p localize to distinct peripheral compartments (Chang et al. 2005). However, since Atg2p requires Atg9p for its localization, we suggest that these compartments may transiently interact. Ba sed on our data, it is possible that Sar1p may influence the assembly of both PC2 and PC9 compartments. Indeed, the structure of the PC2 compartment was altered and there appeared to be more PC9 foci present in cells expressing the Sar1p mutants. Therefore, we next exam ined whether Atg2p and Atg9p colocalize in the presence of the Sar1p mutant proteins. We coexpressed RFP-Atg9p and GFP-Atg2p in WDY64 and WDY65 cell lines. In the absence or pres ence of copper, RFP-Atg9p was found within the lumens of the vacuole and SM (Fig. 6-3). We reported this observation previously and suggested that the lumenal RFP was the result of N-te rminal proteolysis while Atg9p was at these membranes (Chang et al. 2005). Furthermore, despite th e expression of Sar1pT34N or Sar1pH79G, Atg9p did not colocalize with Atg2p (Fig. 6-3). 156

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Finally, we examined the cellular distribu tion of GFP-Atg8p in glucose-adapted cells expressing the mutant forms of Sar1p (Fig. 64). Under normal conditions, Atg8p localizes to the PAS from which it transits to the newly assembled MIPA. At 2h of glucose adaptation without copper, GFP-Atg8p was present throughout the cytosol and localized specifically to one or two MIPAs situated between the adjoini ng sequestering membranes labeled with FM4-64 (Fig. 6-4, white arrows). When the expressi on of Sar1pT34N or Sar1pH79G is enhanced by copper, GFP-Atg8p was observed in multiple punctat e (Fig. 6-4, white arrowheads) and flattened MIPA-like structures (Fig. 6-4, ye llow arrows). However, these structures were not associated with the sequestering membranes, but randomly distributed throughout the cell. Our data suggest that in the presence of Sar1pT34N or Sar1pH79G, MIPA fails to assemble, thereby inhibiting the expansion a nd completion events. We also noticed that the PAS, which normally appears as one or two foci, was structurally different when Sar 1pT34N or Sar1pH79G were expressed. Therefore, we examined the cellular distribution of GFPAtg17p, a structural component of the PAS not present at the MIPA (Fig. 6-12). During glucose adaptation in the absence of c opper, GFP-Atg17p was found at one or two distinct foci juxtaposed to the vacuole or sequestering membranes. However, in the presence of copper, GFP-Atg17p was present in foci and larger diffuse structures of irregular shapes that were distant from the vacuole and SM. The data suggest that Sar1p may play a role in the maintenance of the PAS. The Lipidation Of Atg8p During Gluco se-Induced Micropexophagy Is Altered Upon Expression Of Dominant-Negative Sar1p Mutants Based on GFP-Atg8p labeling, we have determined that the MIPA fails to assemble when either Sar1pT34N or Sar1pH79G are expressed. Atg8p not only localizes to the MIPA, but is essential for its formation. Upon proteolytic activation by Atg4p, the C-terminal glycine of 157

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Atg8p becomes conjugated by a thio-ester linkage to cysteine 518 of Atg7p. Atg8p is then conjugated to phosphatidylethanolamine (PtdEtn) by the actions of Atg3p. We have previously shown that Atg7p-HA forms a DTT-se nsitive thio-ester conjugate w ith a protein whose size we estimated at 25 kDa (Yuan et al., 1999). When the active cysteine of Atg7p is mutated to a serine (Atg7pC518S-HA) this conjugation be comes DTT resistant. Therefore, when Atg7pC518S-HA is expressed in WT cells, we ob serve two protein bands at 70 kDa and 95 kDa (Fig. 6-5A). The upper band was not observed in cells lacking Atg8p sugge sting that the 95 kDa band represented Atg7pC518S-HA c onjugated by an ester linkage to Atg8p. In order to assess the effects of Sar1pT34N or Sar1pH79G on Atg7p activity, we expressed Atg7pC518S-HA in these mutants. These cells were grown in YNM and switched to YND for 3h and 6h with and without copper sulfate. Afterwards, the prot ein conjugate of Atg7p a nd Atg8p was visualized by Western blotting (Fig. 6-5A). In the absence and presence of copper, both unconjugated (70 kDa) and conjugated (95 kDa) forms of Atg7pC 518S-HA were observed suggesting that Atg7p activity was not affected by eith er Sar1pT34N or Sar1pH79G. We next examined the effects of Sar1pT34N or Sar1pH79G on the lipidation of Atg8p. This was done by examining the processing of At g8p on Western blots given that the lipidated form of Atg8p migrates at a lower molecular mass on SDS-PAGE (Stromhaug et al., 2004). Cells were grown in YNM and switched to YND with and without copper sulfate for 0 9h. At 0h, GFP-Atg8p migrated as a single band around 40 kDa (Fig. 6-5B). At 3 9h in the absence of copper, smaller forms of GFP-Atg8p began to a ppear with the lipidated form migrating at 35 kDa. When copper was present, the 35 kDa protein was virtually ab sent. The data suggest that the expression of Sar1pT34N or Sar1pH 79G altered the lipidation of Atg8p. 158

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Sar1p Is Required For Early And Late Events Of Macropexophagy We have shown that Sar1pT34N and Sar1pH79G suppress the de gradation of AOX during ethanol-induced macrope xophagy. During macropexophagy, individual peroxisomes are engulfed by sequestering membranes that can be visualized by GFP-At g8p. Like the MIPA, these membranes are presumed to be organized from the PAS (Dunn et al., 2005). In cells not expressing Sar1pT34N and Sar1pH79G, GFP-Atg8p was present at the PAS (Fig. 6-6, yellow arrowheads), pexophagosomes (Fig. 6-6, white arrows), and within the vacuole (Fig. 6-6A). We next examined the effects of Sar1pT34N and Sar1pH79G on the assembly of pexophagosomes as visualized by GFP-Atg8p labeling (Fig. 6-6) and ultrastructural morphology (Fig. 6-7). When Sar1pT34N is expressed, peroxisomes were not sequestered within pexophagosomes or the vacuole, but observed in the cytosol. GFP-Atg8 localized to the PAS (Fig. 6-6, yellow arrowhead) and crescent-like structures (Fig. 67B), which may represent the nucleation of sequestering membranes that we observed near th e peroxisomes (Fig. 6-7B, arrow). In cells expressing Sar1pH79G, we observed pexophagosomes with foci containing GFP-Atg8 (Fig. 66D, white arrows), but the diffuse vacuole labelin g consistent with vacuolar fusion was absent. Ultrastructural analyses revealed that th ese pexophagic vacuoles were bound by multiple membranes (Fig. 6-7C, arrowheads). Furtherm ore, intact peroxisomes bound by a single membrane could be observed within the vac uole (Fig. 6-7D, arrowh eads). RFP-Atg9p was found within the vacuole or localized to the vac uole surface and foci juxtaposed to the vacuole (Fig. 6-6A and 6-6C). This distribution was unaff ected by the expression of either Sar1p mutant (Fig. 6-6B and 6-6D). RFP-Atg9 could be observed with some G FP-Atg8 foci at the pexophagosome (Fig. 6-7, white arrow). These obs ervations combined with our biochemical data (Fig. 6-1) suggest that the pexophagosomes present in cells expressing Sar1pH79G may fuse poorly with the vacuole or are resistant to ly tic enzymes of the vacuole. We have shown that 159

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Sar1pT34N suppresses the formation of th e pexophagosome, while the delivery of the peroxisome from the pexophagosome to the vacuole for degradation appears to be defective in cells expressing Sar1pH79G. RFP-HDEL Localizes To Pexophagosomes When Sar1pH79G Is Expressed These dominant-negative Sar1p mutants have been shown to effectively alter the trafficking of ER proteins in S. cerevisiae (Ward et al., 2001). Since the ER has been implicated in the formation of autophagic vacuoles in mammalian cells (Dunn, 1990), we decided to examine the localization of RFP-HDEL during macropexophagy (Fig. 6-8). RFP-HDEL with its amino-terminal signal sequence and carboxy-termin al HDEL retrieval signal is targeted to the ER lumen by the receptor Erd2p whic h binds the HDEL sequence (Bevis et al. 2002). Upon ethanol adaptation, RFP-HDEL was found exclusively at the ER located adjacent to the cell periphery and around the nucleus (Figs. 6-8 A and D). The distribution of RFP-HDEL was unaltered in ethanol-adapted cells expressing Sar1pT34N (F igs. 6-8 B and C). When Sar1pH79G is expressed during 2h of ethanol adaptation, we obs erved RFP-HDEL at the ER and engulfing individual peroxisomes (Fig. 6-8, arro ws). At times longer than 3h, the RFP was observed within the vacuole (Fig. 68, arrowheads). These pexophagosomes did not contain Sec7p, a protein of the Golgi apparatus (unpublis hed observations). The results suggest that components of the ER are observed with the pex ophagosomes when ER recycling is inhibited by Sar1pH79G. Discussion Earlier reports have demonstrated that some Sec proteins are required for autophagy in S. cerevisiae. Three of these proteins, ScSec12p, ScSec23p, and ScSec24p, are essential for ScSar1p activity and trafficking of proteins out of and into the ER. We examined the role of Sar1p in the selective degr adation of peroxisomes by pe xophagy by tightly regulating the 160

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expression of two dominant-nega tive mutants of Sar1p. Sar1pT34N has a high affinity for GDP and suppresses COPII-dependent protein exit from the ER. Sar1pH79G has a defective GTPase thereby inhibiting the recycling of proteins back to the ER. We have shown that this protein is essential for starvation-induced autophagy, glucose-induced micr opexophagy, and ethanolinduced macropexophagy. How is it possible that Sar1p, which is essential for trafficking proteins between the ER and the Golgi, ha s such a far-reaching influence on microand macropexophagy? In this study, we attempted to a ddress this question to define the roles of Sar1p in these diverse autophagic pathways. Role of Sar1p in Micropexophagy Our data suggest that the de gradation of AOX is dramatically reduced when either Sar1pT34N or Sar1pH79G are expressed. The pe roxisomes were observed almost completely engulfed by sequestering membranes out side the vacuole. This sugge sts that the blockage occurs at a late sequestration event (e .g., expansion and/or completion). Moreover, MIPA formation did not occur when either Sar1pT34N or Sar1pH79G were expressed. The MIPA is a crescentshaped structure that joins the apposing sequestering membranes to initiate their fusion thereby completing expansion and completion events. Th is structure contains Atg8p and Atg26p and its formation requires a number of Atg proteins including Atg9p and Atg11p. Therefore, we examined the trafficking of Atg9p and Atg11p a nd showed that their trafficking to the sequestering membranes was unaffect ed by the Sar1p mutants. The MIPA assembles from the PAS or nucle ation complex, a structure that contains Atg8p and Atg17p (Oku et al., 2006; Yamashita et al., 2006). The formation of the MIPA is dependent upon the lipidation of Atg8p, wh ich requires Atg4p, Atg7p, and Atg3p (Mukaiyama et al. 2004). Our data suggest that the inability of cells expressing the Sar1p mutants to assemble the MIPA may be related to their inabili ty to lipidate Atg8p (see Fig. 6-5B). Atg4p 161

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proteolytically activates At g8p. Atg7p forms a high-energy conjugate with Atg8p thereby delivering Atg8p to Atg3p for its conjugation to p hosphatidylethanolamine. The data show that the formation of the high-energy conjugate w ith Atg7p proceeds normally when these Sar1p mutants are expressed (see Fig. 6-5A ). Therefore, in inability to lipidate Atg8p may be related to an inactivation of Atg3p or a decrease in the av ailability of PtdEtn. The effects of Sar1p on Atg3p activity are unknown and remain to be examine d. However, the availability of PtdEtn has been shown to be essential fo r the autophagic response (Nebauer et al. 2007). Thus, the inability to form the MIPA ma y be related to lower levels or missorting of PtdEtn. The availability of PtdEtn was reduced in cells l acking ScVps4p or ScVps36p, proteins required for post-Golgi trafficking. The autophagy-like transp ort of cytosolic pAPI to the vacuole was inhibited in these mutants (Nebauer et al. 2007). Meanwhile, ethanolamine rescued pAPI transport, suggesting that PtdEtn was a limiting fact or. Furthermore, mutants defective in PtdEtn production show reduced ScAtg8p localization to the PAS and suppressed Cvt and autophagy pathways (Nebauer et al. 2007). These effects were not due to reduced ScAtg8p levels, but to low levels of PtdEtn, suggesting th at the availability of PtdEtn ma y be critical to the lipidation of Atg8p and assembly of the PAS. Psd2p, which s ynthesizes PtdEtn from phosphatidylserine, can interact with Sec28, a protein of the COPII complex (Schuldiner et al. 2005). The trafficking of lipids out of the ER is thought to be mediated by both vesi cular and non-vesicular traffic (Levine, 2004). However, the movements of Pt dEtn to the cell surfac e were not affected by brefeldin A and thus, may not require Sar1p (Vance et al. 1991). Nevertheless, the effects of Sar1p on the cellular levels of Pt dEtn remain to be studied. Atg26p is also required for MIPA forma tion. Phosphatidylinositol 4-monophosphate (PI4P) transits from the ER to the PAS or nucleation complex thereby recruiting Atg26p by 162

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interacting with its GRAM domain (Yamashita et al., 2006). The sterol conversion of Atg26p is essential for the formation of the MIPA. It is pos sible that the trafficking of PI4P may be altered in cells expressing the Sar1p mutants. Indeed, Sar1p has been implicated in the exit of glycosylphosphatidylinositol (GPI) anchored G FP from the ER (Stephens and Pepperkok, 2004). However, we have no data to suggest that PI4P transport is influenced by Sar1p, and studies are currently ongoing. In summary, the expression of Sar1pT34N or Sar1pH79G inhibits glucose-induced micropexophagy by suppressing the assembly of the MI PA. The most probable scenario is that Sar1p affects Atg8p lipidation an d/or the trafficking of Atg26p which is essential to MIPA formation. The suppressed lipidation of Atg8 may be due to an inactivation of Atg3p or insufficient PtdEtn, while the movements of Atg26p require vesicular traffi cking of PI4P from the ER to the PAS. Role of Sar1p in Macropexophagy The data from sec mutants provides circumstantial evidence that the autophagosome membranes may originate from the ER (Ishihara et al. 2001; Hamasaki et al. 2003; Reggiori et al. 2004). In this study, we characterized the essential role of Sar1p in macropexophagy in Pichia pastoris Ethanol-induced peroxisome degradat ion is suppressed when cells express either Sar1pT34N or Sar1pH79G (Fig. 6-1). However, these proteins inhibit different events of macropexophagy. We have shown that pexophagos omes do not form when Sar1pT34N is expressed. ScSec12p, a GDP/GTP exchange factor that influences ScSar1p activity, is also required for the formation of autopha gic vacuoles in nitrogen-starved S. cerevisiae (Ishihara et al. 2001; Reggiori et al. 2004). Brucella abortus is an intracellular pathogen that infiltrates the autophagosome to replicate in vacuoles that contain the ER protein calnexin (Celli et al. 2005). When HsSar1pT39N is expressed, this bacterium fa ils to replicate or localize with calnexin163

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positive autophagic vacuoles, but instead is f ound in LAMP1-positive phagolysosomes (Celli et al. 2005). These results subs tantiate our findings that HsSar1pT39N and its P. pastoris analogue Sar1pT34N suppresses autophagic vacuole formation. In the case of B. abortus since the replication vacuole is not fo rmed, this bacterium transits to the phagolysosome where it is killed and degraded. On the other hand, bacteria l replication with in calnexin-positive autophagic vacuoles was not altered by Sar1pH79G. This is consistent with our findings that Sar1pH79G does not affect formation of the pexophagosome. Pexophagosomes with GFP-Atg8p at their delimiting membranes can be observed in cells expressing Sar1pH79G. Their morphology is cons istent with fully formed pexophagosomes (see Fig. 6-7C). Meanwhile, profiles of pexophagosomes delimited by GFP-Atg8 can be occasionally observed within the vacuole (see Figs. 6-6C and 6-7D). The data suggest that Sar1pH79G may interrupt fusion of the pexophagosom e with the vacuole and/or degradation of the pexophagosome within the vacuole. It is possible that the trafficking to the vacuole of the fusion machinery (Ypt7, Vti1, and Vam3) or th e degradative lipases (A tg15) or proteinases (Pep4 and Prb1) may have been altered by Sar1pH79G (Levine and Klionsky, 2004). On the other hand, alterations in the membranes of th e pexophagosome may interrupt fusion and lysis. Indeed, we have observed that the delimiting membranes of the pexophagosomes also contained RFP-HDEL, a marker for the ER membrane pr otein Erd2 (HDEL receptor), an ER membrane protein. At later times of macropexophagy, the RFP was found within the vacuole suggesting that there was some delivery of the pexophagosome to the vacuole. At no time in wild type cells did we observe RFP-HDEL delimiting the membranes of the pexophagosome or within the vacuole. These results are consistent with Sa r1pH79G inhibiting retrogra de trafficking of the RFP-HDEL and other ER compone nts from the pexophagosome to the ER. The presence of 164

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these ER components may ultimately suppre ss pexophagosome fusion with the vacuole and provide resistance to vacuolar lipas es and proteinases. Thus, our data suggest that recycling of ER components from the pexophagosome is criti cal for delivery of the peroxisomes to the vacuole for degradation. In summary, we propose that the movement of ER components into and out of the pexophagosome is influenced by Sar1p (Fig. 6-9). We have utilized dominant negative mutants of Sar1p that inhibit anter ograde (Sar1pT34N) and retrograde (Sar1pH79G) movements of ER proteins. We have shown that anterograde move ments of ER proteins to the pexophagosome are critical for its formation, while retrograde m ovements of ER proteins from the pexophagosome are essential for the delivery of the peroxisome to the vacuole. Summary We have utilized Sar1p mutants defective in anterograde (Sar1pT34N) and retrograde (Sar1pH79G) transport of ER proteins to demonstr ate that Sar1p is essential for autophagy and pexophagy. These mutants suppress micropexophagy by inhibiting the formation of the MIPA and delivery of the peroxisomes to the vacuol e. In regards to macropexophagy, inhibition of anterograde ER transport suppresses the formation of the pexophagosome. Meanwhile, inhibition of retrograde ER transport does not affect pexophagosome formation, but instead suppresses pexophagosome fusion with vacuole and/ or degradation by the vacuolar enzymes. Our results demonstrate that pr otein sorting and vesicular m ovements directed by Sar1p are essential for microand macropexophagy. 165

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Table6-1: Pichia pastoris strains Name Genotype Reference GS115 his4 (Cregg et al. 1985) PPF1 arg4 his4 (Yuan et al. 1997) DMM1 GS115::pDM1 (PAOX1BFP-SKL, ZeocinR) (Kim et al. 2001) AJM38 GS115 his4 ::pWD18 (PATG7 atg7 -HA (C518S), HIS4 ) This study WDY64 PPF1 his4 arg4::pWD16-Sar1(T34N) (PCUP sar1 (T34N), ARG4 ) This study WDY65 PPF1 his4 arg4::pWD16-Sar1(H79G) (PCUP sar1 (H79G), ARG4 ) This study WDY66 WDY64 his4 ::pWD21 (PGAPDH GFPATG8 HIS4) This study WDY67 WDY65 his4 :: pWD21 (PGAPDH GFPATG8 HIS4) This study WDY70 DMM1 his4 ::pWD21 (PGAPDH GFPATG8 HIS4) This study WDY111 SJCF257 arg4 his4 atg8::pWD18(PATG7 atg7 -HA (C518S), HIS4 ) This study WDY120 LAM44 :: pAJM6 (PGAPDH mRFPATG9 ZeocinR) This study WDY121 LAM45 :: pAJM6 (PGAPDH mRFPATG9 ZeocinR) This study WDY122 WDY66 :: pAJM6 (PGAPDH mRFPATG9 ZeocinR) This study WDY123 WDY67 :: pAJM6 (PGAPDH mRFPATG9 ZeocinR) This study WDY126 WDY64 his4 ::pWD18(PATG7 atg7 -HA (C518S), HIS4 ) This study WDY127 WDY65 his4 ::pWD18(PATG7 atg7 -HA (C518S), HIS4 ) This study LAM13 DMM1 his4 ::pSar1(T34N) (PCUP sar1 (T34N), HIS4) This study LAM14 DMM1 his4 ::pSar1(H79G) (PCUP sar1 (H79G), HIS4) This study LAM40 WDY64 his4 :: pTC1 (PGAPDH GFPATG9 HIS4) This study LAM41 WDY65 his4 :: pTC1 (PGAPDH GFPATG9 HIS4) This study LAM42 WDY64 his4 ::pPS64 (PGAPDH GFP/HAATG11, HIS4 ) This study LAM43 WDY65 his4 ::pPS64 (PGAPDH GFP/HAATG11, HIS4 ) This study LAM44 WDY64 his4 ::pPS69 (PGAPDH GFPATG2 HIS4) This study LAM45 WDY65 his4 ::pPS64 (PGAPDH GFPATG2 HIS4) This study LAM46 WDY64 his4 ::pPS55-G12 (PGAPDH GFP/HA-ATG18, HIS4) This study LAM47 WDY65 his4 ::pPS55-G12 (PGAPDH GFP/HA-ATG18, HIS4) This study LAM54 WDY64 his4 ::pWD24 (PGAPDH GFPATG17 HIS4) This study LAM55 WDY65 his4 ::pWD24 (PGAPDH GFPATG17 HIS4) This study LAM63 WDY64 his4 ::pIB2-dsRED-HDEL (PGAPDH dsRFP-HDEL, HIS4) This study LAM64 WDY65 his4 ::pIB2-dsRED-HDEL (PGAPDH dsRFP-HDEL, HIS4) This study 166

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Figure 6-1. Sar1p is essential for autophagy and pexophagy. Autophagy (A), glucose-induced micropexophagy (B) and ethanol-induced macropexophagy (C) were evaluated in wild-type cells and cells expressing Sar1pT34N and Sar1pH79G. In panel A, endogenous proteins of GS115, WDY64, and WDY65 cells were labeled with 14C-valine for 16 hours in YND. The cells were then washed and switched to SD-N medium ( CuSO4) supplemented with 10 mM valine. Aliquots were removed at 2-24 hours of chase and precipitated with TCA. The rates of protein degradation based on the produc tion of TCA-soluble radioact ivity were expressed as a percentage relative to untreated controls. In panel B, GS115, WDY64, and WDY65 cells were grown in YNM and then switched to YND ( CuSO4) at which time the loss of AOX activity was measured over 6h. The data represents the mean SD of 4-6 trials. In panel C, GS115, WDY64, and WDY65 cells were grown in YNM and then switched to YNE ( CuSO4). At 024h samples were prepared for SDS-PAGE and the remaining AOX visualized by Western blotting. The data represents a representative blot of 3 different trials. 167

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Figure 6-2. The expression of Sar1pT34N or Sar 1pH79G suppresses a late sequestration event of micropexophagy. WDY64 (A, B), WDY65 (E and F), LAM13 (C, D), and LAM14 (G, H) were adapted from YNM to YND in the absence (A, E) and presence (B, C, D, F, G, H) of CuSO4 for 2-3 hours. Cells were then fixed and prepared fo r viewing by electron micros copy (A, B, E, F) or viewed in situ by fluorescence microscopy (C, D, G, H). In cells not treated with CuSO4, the peroxisomes (P) could be found within the va cuole (A) or surrounded by membranes derived from the vacuole (E). When cells expre ssed either Sar1pT34N (B) or Sar1pH79G, the peroxisomes were partially surrounded by the vacuole and sequestering membranes, but not observed within the vacuole. An extensive, ribbon-like endoplasmi c reticulum (ER) was observed in cells expressing Sar1pT34N while a multilamellar Golgi (Go) apparatus was present with Sar1pH79G expression. By fluorescence microscopy, the vacuole and those sequestering membranes derived from the vacuole were observed by staining with FM4-64, and the peroxisomes were visualized by expressing BFP-SK L. In cells expressing either Sar1pT34N or Sar1pH79G, the peroxisomes were found to be almost completely surrounded by sequestering membranes. These membranes stained with FM464 and appeared as vesiculated extensions of the vacuole. 168

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Figure 6-3. The trafficki ng of Atg11p, Atg2p and Atg9p during glucose-induced micropexophagy is not dependent upon Sar1p. WDY64 and WDY65 cells expressing GFPAtg11p, GFP-Atg2p, GFP-Atg9p or GFP-Atg2p and RFP-Atg9p were adapted from YNM to YND in the absence and presence of CuSO4. At 2h of glucose adap tation, the cells were visualized by fluorescence microscopy. The cellular localizations of Atg11p, Atg2p and Atg9p were unaltered in cells expressing Sar1pT34N or Sar1pH79G. GFP-Atg11p was present at the vacuole and sequestering membrane s. GFP-Atg2p was visualized at peripheral foci. However, in the presence of the Sar1p mutants, these foci appeared to be greater in number and sometimes coalesced forming irregular structures (yellow arrows). GFP-Atg9p was observed at the vacuole and sequestering membranes labeled with FM464. Atg9p could also be visualized at perivacuolar structures (white arrows), peripheral foci (white arrowheads), and the PAS at the tips of the sequestering membrane s (yellow arrowheads). The coalescing foci of Atg2p did not contain RFP-Atg9p. 169

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Figure 6-4. Sar1p is essential for the formati on of the MIPA. WDY 64 and WDY65 cell lines expressing GFP-Atg8 were adapted from YNM to YND in the absence and presence of CuSO4. At 2h of glucose adaptation, the cells were visualized by fluorescence microscopy. The vacuole and those sequestering membrane s derived from the vacuole were observed by staining with FM4-64 while the MIPA was observed as a crescent-shaped structure labeled with GFP-Atg8 (arrows). In the absence of copper, th e MIPA was found bridging the FM4-64 labeled membranes. However, when the expression of either Sar1pT34N or Sar1pH79G was induced with CuSO4, GFP-Atg8p localized to one or more dots that appeared to be distributed throughout the cell (arrowheads). Occasionally, GFP-Atg8p could be found in a flattened MIPA-like structure, but not situated near sequ estering membranes (yellow arrows). 170

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Figure 6-5. The effects of Sar1p on the lipida tion of Atg8p. In panel A, Atg7pC518S-HA was expressed in GS115, SJCF257 ( atg8), WDY64 and WDY65 cells. The cells were adapted from YNM to YND in the absence and presence of CuSO4 for 3h (left panel) and 6h (right panel). At that time, the cells were solubilized in SDS sample buffer, and the 75kDa Atg7p and 100kDa Atg7p protein-protein conj ugate visualized by Western blotti ng. The protein-protein conjugates of Atg7pC518S were observed in wild-type cel ls and in cells expressing Sar1pT34N or Sar1pH79G, but not in cells l acking Atg8p. In panel B, WDY 64 and WDY65 cells expressing GFP-Atg8p were adapted from YNM to YND in the absence and presence of CuSO4 for 0-9h. The cells were then solubilized and the proces sing of Atg8p examined by Western blot. During glucose adaptation, a 38kDa form of GFP-Atg8p was observed at 3-6h. This protein was not present when Sar1pT34N or Sar1pH79G was expre ssed. The slower migration of this protein compared the GFP-Atg8p is consistent w ith the lipidated form of Atg8p. 171

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Figure 6-6. Effects of Sar1pT34N and Sar1pH79G on ethanol-induced macropexophagy. WDY64 (A and B) and WDY65 (C and D) cell lines expressing GFP-Atg8p and RFP-Atg9p were adapted from YNM to YNE in the absence (A and C) and presence (B and D) of CuSO4. At 2h of ethanol adaptation, the cells were vi sualized by fluorescence microscopy. In the absence of CuSO4, GFP-Atg8p was found at the PAS (yellow arrowheads) and pexophagosomes (white arrows). RFP-Atg9p was at foci juxt aposed to the vacuole or the pexophagosome. Colocalization of RFP-Atg9p and GFP-Atg8p at th e PAS or pexophagosome was rare, but both proteins could be observed within the vacuole. When Sar1pT34N is expressed, GFP-Atg8p was found almost exclusively at foci presumed to be the PAS. GFP-Atg9p was observed within the vacuole and at perivacuolar foci that lacked GFP-Atg8p. To th e contrary, when Sar1pH79G is expressed, GFP-Atg8p was found at the PAS and pexophagosome. GFP-Atg9p was visualized at perivacuolar foci and within the vacuole. 172

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Figure 6-7. Sar1pT34N suppresses formati on of the pexophagosome while Sar1pH79G suppresses delivery of the peroxisomes from the pexophagosome to the vacuole. WDY64 and WDY65 cell lines expressing GFPAtg8p were adapted from YNM to YNE in the presence of copper sulfate. At 2h of ethanol adaptati on, the cells were visualized by fluorescence microscopy or fixed and processed for electr on microscopy. When Sar1pT34N was expressed, GFP-Atg8p was found at the PAS (A) and occasiona lly at crescent-shaped structures (B). Indeed, membranes were occasionally observed near peroxisomes, but not engulfing them (black arrow). Meanwhile, in cells expressing Sar1pH79G, pexophagosomes containing GFP-Atg8p were observed (C and D). Distinct foci containing GFP-Atg8p and RFP-Atg9p (white arrows) were observed at the pexophagosome. The pexophagosome bound by multiple membranes was visualized by electron microscopy (C, arrowheads). At times, we also observed remnants of GFP-Atg8p membranes and single membrane-bound undegraded peroxisomes within the vacuole (D, arrowheads). Trafficking of R FP-Atg9p to the vacuole was unaltered by the expression of either Sar1p mutant. N, nuc leus; P, peroxisome; and V, vacuole. 173

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Figure 6-8. Components of the endoplasmic reticulum are present at the pexophagosome when cells express Sar1pH79G. WDY64 (A, B, and C) and WDY65 (D, E, and F) cell lines expressing RFP-HDEL were adapted from YNM to YNE in the absence (A and D) and presence (B, C, E, and F) of CuSO4. At 2h (A, B, D, and E) and 3h (C and F) of ethanol adaptation, the cells were visualized by fluorescence microscopy. RFP-HDEL wa s localized solely to the ER situated near the cell periphery and around the nucleus. This distri bution was unaltered during macropexophagy (A and D). When Sar1pT34N was expressed, the cellular distribution of RFPHDEL was restricted to the ER (B and C). However, at 2h of ethanol adap tation in the presence of Sar1pH79G, RFP-HDEL was visualized not only at the ER but also at vacuoles containing phase-dense peroxisomes (E, white arrows). At 3h, RFP-HDEL could be detected within the yeast vacuole (F, white arrowheads). 174

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Figure 6-9. Model of the role of Sar1p in macropexophagy. During ethanol adaptation, peroxisomes (P) are engulfed by pexophagosomes that are assembled from sequestering membranes (SM) and expanded by the PAS with membranes containing Atg8p. Based on our data, we propose that Sar1p is required for the movements of ER proteins into and out of the pexophagosome. The pexophagosome assembles with the assistance of Sec13p and Sar1p delivery of ER proteins. The anterograde transport of ER proteins to the forming pexophagosome is inhibited by Sar1 pT34N which is unable to exchange GTP for GDP. Once the pexophagosome is assembled, the ER proteins ar e recycled back to th e ER with the aid of Sar1p. Retrograde transport of the ER proteins from the pexophagosome is inhibited by Sar1pH79G which is unable to hydrolyze GTP. 175

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Figure 6-10. Effects of Sar1pT34N and Sar1pH 79G on cell growth. Sar1pT34N and Sar1pH79G were grown in minimal medium for 24 hour s in the absence or presence of CuSO4 and growth of the culture assessed by measuring absorbance at 600nm. The effects of the Sar1p mutants on cell growth were minimal at five hours post copp er induction. However, at longer times, growth was dramatically reduced compar ed to untreated cells. 176

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Figure 6-11. Stages of glucose-induced micropexophagy. Cells expre ssing BFP-SKL were adapted from YNM to YND for 2h. Cells at all stages of micropexophagy can be visualized by uorescence microscopy. During YNM growth, the peroxisomes visualized by BFP-SKL were situated next to a round vacuole visualized by FM4-64. During nucleation, the sequestering membranes that arise from the vacuole only partially extend around the peroxisomes. The expansion stage was characterized by sequest ering membranes extending 50 to 90% around the peroxisomes. The completion stage was characterized by peroxisomes fully surrounded by sequestering membranes that appear segmented and not fused. Finally, the degradative stage was characterized by continuous FM4-64 labeled membranes enclosing peroxisomes that were not clearly delineated. fl 177

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micropexophagy in cells expressing Sar1pT34N or Sar1pH79G. WDY6 4 and WDY65 c At 2h of glucose adaptation, the cells were visualized by fluorescence microscopy. Regardle sequestering membranes. In untreated cells, G FP-Atg17p was visualized at 1-3 foci (PAS) near foci that at times appeared to coalesce (yellow arrows). Figure 6-12. Cellular localization of Atg18p and Atg17p during glucose-induced ells expressing GFP-Atg18p (A) or GFP-Atg17p (B) were adapted from YNM to YND ( CuSO4). ss of the presence of the Sar1p mutants, GFP-Atg18p trafficked normally to the vacuole and the vacuole. In cells expressing Sar1pT34N or Sar1pH79G, GFP-Atg17p localized to multiple 178

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CHAPTER 7 CONCLUSION These studies first explain how to procure a restriction enzyme-mediated integration mutant (REMI) (Chapter 2). Using this applicati on, we have identified a number of ATG genes essential for pexophagy and autophagy. In Chapter 3, I characterized one such gene, ATG9 that encodes an integral membrane protein with 5-6 transmembrane domains. At this time, Atg9 was the only known transmembrane protein required for pexophagy. We next evaluated the s ubcellular localization of Atg9 by tagging GFP to its N-terminus. In fed cells, I showed that Atg9 localized to a unique peripheral compartment (PC9) that did not contai n protein markers for endoplasmic reticulum, transition ER, Golgi apparatus, or mitochondr ia. Upon glucose-indu ced micropexophagy, Atg9 was found at perivacuolar structures (PVS) situat ed at the base of the sequestering membranes (SM) that arose from the vacuole and at the SM. Because Atg9 was present in multiple compartments during pexophagy, my aim was to define its trafficking in the hopes of better understanding the assembly and expansion of the SM during pexophagy. I also showed that the trafficking of Atg9 to the PVS required Vps15 and Atg11. Movements of Atg9 from the PVS to the SM required Atg2 and Atg7. These studies were the first to demonstrate the movements of a membrane protein to the expandi ng sequestering membranes. In Chapters 4 and 5, I set out to define the functional motifs within Atg9. Atg9 is present throughout all eukaryotes, and its homologous regions were examined by FASTA analysis. Atg9 membrane motifs of import were predicted by in situ analysis to increase the probability that my site-directed mutagenesis would be productive and efficient. Multiple motifs were shown to be important for Atg9 function and tr afficking. I have shown that Atg9 has two domains required for ER exit (one of which is ch aracterized by two glutamates), one di-lysine-like motif required 179

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180 ophagy ER in pexophagy. Unfortunately, the very early even ts regulating pexophagy are poorly understood. Moreover, there has been considerable heated discussion concerning the origin of the pexophagic membranes. Whether the ER contributes to th ese membranes or the membranes are synthesized de novo has been a topic of heated debate for twenty years. Sar1 is the GTPase responsible for the generation of COPII vesicles. Thus, the study of dominant-negative Sar1 mutants in regards to the trafficking of Atg protei ns appeared to be a straight forward approach to addressing pexophagic regulation and the forma tion of pexophagosomes. Sar1 appears to be the earliest post-translational protein factor influencing pexophagy. The findi ng that Sar1 is required for micropexophagic-specific membrane apparatus (MIP A) formation suggests a requirement for COPII trafficking in SM expansion and comple tion during micropexophagy. I also showed that cells expressing sar1T34N, which suppresses protein exit from the ER, cannot form autopexophagosomes. Meanwhile, cells expressi ng sar1pH79G, which inhibits retrograde transport back to the ER, do form autopexophagosomes. These autopexophagosomes contain an ER marker protein suggesting that sar1H79G i nhibits the recycling of the ER proteins. I am the first to characte rize the roles of Atg9 and Sar1 in the expansion of the sequestering membranes during micropexophagy. Thes e studies have provided new directions and molecular tools to examine these ev ents during macropexopha gy and autophagy. for PC9 exit, one motif necessary for its asso ciation with the SM and may associate with peroxisomal membrane proteins, and one domain not required for trafficking to the SM but is essential for Atg9 function. A ll of these motifs are required for Atg9 function during pex and autophagy, suggesting that Atg9 functions at the SM. In Chapter 5, I showed that Sar1p is essential for pexophagy and autophagy. Our expanding knowledge of the field of pexophagy has le d to the pursuit of th e role of the

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BIOGRAPHICAL SKETCH Laura Aaron (Miller) Schroder graduated from Saint John Lutheran School in Ocala, FL in 1997. She earned an Associate in Arts from Central Florida Community College in Ocala, FL in 1999 and was inspired by Lowell Sanders, Ph.D. to pursue a Bachelor of Science in Chemistry from the University of South Florida in Tampa, FL. Lauras work in the laboratory of David J. Merkler, Ph.D. on glutathione as subs trate for the enzyme peptidylglycine -amidating monooxygenase cumulated in a poster at the Amer ican Society for Biochemistry and Molecular Biology conference and her first fi rst-author paper. She was s upported during this time with a summer research stipend from the Institute for Biomolecular Science during which she was mentored by Clark Craddock, Ph.D., J.D. La ura was also an undergraduate and 3 month postbaccaulaureate researcher in the la b of Denise R. Cooper, Ph.D. on PKCII alternative splicing. Laura began her graduate work in the Interdisciplinary Pr ogram in Biomedical Sciences, part of the College of Medicine at th e University of Florida in 2001 August. Laura joined the lab of William A. Dunn, Jr. in 2002 December upon the recommendation of John Aris, Ph.D. and has authored three papers since then. The test of a first-rate intelligence is the ability to hold two opposed idea s in mind at the same time, and still retain the ability to function. F. Scott Fitzgerald in The Crack Up 190