Mitochondrial-localized

Phosphatidylethanolamine in Mitochondrial Dynamics and Autophagy

by

Eliana Y.L. Chan

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Graduate Department of Biochemistry University of Toronto

© Copyright by Eliana Y. L. Chan 2014

ii

Mitochondrial-localized Phosphatidylethanolamine in

Mitochondrial Dynamics and Autophagy

Eliana Y. L. Chan

Doctor of Philosophy

Graduate Department of Biochemistry University of Toronto

2014

ABSTRACT

Phosphatidylethanolamine (PE) constitutes a significant proportion of total phospholipids

in biological membranes. A major route of PE biosynthesis occurs via the decarboxylation of

phosphatidylserine (PS) by the inner-mitochondrial-membrane (IMM)-localized PS

decarboxylase (PSD). The only other known phospholipids that are synthesized in mitochondria

are the mitochondrial-enriched phosphatidylglycerol (PG) and cardiolipin (CL); the synthesis of

other biological lipids occurs mostly in the endoplasmic reticulum (ER). Hence, the

mitochondrial-localization of PSD led me to speculate that PE within the mitochondrial

membrane might have important functions. In this thesis, I describe my findings that

mitochondrial-localized PE plays an important role in mitochondrial membrane remodelling and

autophagy, a conserved quality control mechanism. Mitochondrial PE promotes mitochondrial

membrane fusion by enhancing the biogenesis of the short isoform of mitochondrial genome

maintenance 1 (s-Mgm1), a key mitochondrial fusion protein. The biophysical properties of PE

on the mitochondrial membrane also promote fusion likely by increasing the rate of lipid mixing

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during fusion. In addition, mitochondrial PE is important for oxidative phosphorylation and the

maintenance of mitochondrial ATP levels, crucial mitochondrial homeostatic functions. I also

demonstrate that a reduction in mitochondrial PE impairs autophagy likely by impeding

autophagosome formation, expansion and/or fusion with the vacuole in yeast. My results

provide additional insight into the functional importance of mitochondrial PE and highlight the

specific lipid composition of the mitochondrial membrane as a key player in mitochondrial

membrane fusion and autophagy.

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ACKNOWLEDGEMENTS

First and foremost, I would like to thank my supervisor, Angus, for his time and patience,

and for giving me the opportunity to learn and grow in his lab. He has been a wonderful mentor

both in and out of the lab and has given me an experience that will be deeply etched in my heart.

I am also immensely grateful to Jeff, our former post-doc, who taught me how to think

critically and make each experiment count. Without Jeff's tutelage, I am sure I will not be the

researcher I am today.

I would also like to thank my supervisory committee members, Dr. Reinhart Reithmeier

and Dr. Grant Brown for their guidance and insightful suggestions throughout my time as a

graduate student.

My gratitude also goes out to all past and present members of the McQuibban lab who

have supported me through the good times and the bad. My experience would not have been the

same without the friendliness and kindness from all members of the lab. I certainly had an

unforgettable time in the McQuibban lab.

I would also like to extend special thanks to Dr. Craig Smibert who has always been

around to give me advice from the time I was an undergraduate student. The members of the

Smibert lab have also been amazing neighbours and have given me invaluable help throughout

the years.

My appreciation also goes out to my wonderful and loving sister who has put up with my

procrastination in booking trips home. I promise I will try to be better!

I would also like to thank Harry and Justine for their continued love, support and

patience, without which, this degree would not have been possible.

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Finally, I would like to dedicate this work to my parents. Their unconditional love and

thoughtful upbringing have made me the person I am today. Despite my shortcomings, they

always believed in me; and for that, I will be eternally grateful.

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TABLE OF CONTENTS

ABSTRACT ...................................................................................................................... ii

ACKNOWLEDGEMENTS ............................................................................................. iv

TABLE OF CONTENTS ................................................................................................. vi

LIST OF FIGURES .......................................................................................................... x

LIST OF ABBREVIATIONS .......................................................................................... xii

CHAPTER 1 INTRODUCTION ..................................................................................... 1

1.1 MITOCHONDRIAL MEMBRANE DYNAMICS ............................................... 1

1.1.1 The discovery of mitochondrial membrane dynamics ................................ 2

1.1.2 Mitochondrial membrane fission ................................................................ 3

1.1.3 Mitochondrial membrane fusion ................................................................. 8

1.1.3.1 Outer mitochondrial membrane fusion .......................................... 8

1.1.3.2 Inner mitochondrial membrane fusion .......................................... 10

1.1.3.3 Coordinating outer and inner mitochondrial membrane fusion .... 13

1.1.3.4 The role of lipids in mitochondrial membrane fusion ................... 15

1.1.4 Mitochondrial membrane dynamics and autophagy ................................... 16

1.2 AUTOPHAGY ...................................................................................................... 17

1.2.1 The autophagy pathway and its molecular machinery ............................... 19

1.2.1.1 The induction of autophagy .......................................................... 19

1.2.1.2 Phagophore formation ................................................................... 21

1.2.1.3 Phagophore expansion and autophagosome formation ................. 21

1.2.1.4 Autophagosome maturation and fusion with the

vacuole/lysosomes ......................................................................... 25

vii

1.2.2 The lipids implicated in autophagy ............................................................. 26

1.2.2.1 Phosphoinositides .......................................................................... 26

1.2.2.2 Phosphatidic acid ............................................................................ 27

1.2.2.3 Membrane composition and/or membrane curvature .................... 28

1.3 PHOSPHATIDYLETHANOLAMINE BIOSYNTHESIS IN

SACCHAROMYCES CEREVISIAE ....................................................................... 30

1.3.1 Phosphatidylserine decarboxylation ........................................................... 30

1.3.2 The Kennedy pathway ................................................................................ 36

1.3.3 Alternative pathways of phosphatidylethanolamine biosynthesis .............. 36

1.3.4 The different contributions of phosphatidylserine decarboxylation and the

Kennedy pathway ....................................................................................... 36

1.3.5 Functions of phosphatidylethanolamine ..................................................... 38

1.4 Specific interests and goals ................................................................................... 39

CHAPTER 2 MATERIALS AND METHODS .............................................................. 41

2.1 REAGENTS .......................................................................................................... 41

2.1.1 Yeast strains ................................................................................................ 41

2.1.2 Plasmids ...................................................................................................... 41

2.1.3 Growth conditions ....................................................................................... 42

2.1.4 Lipids .......................................................................................................... 42

2.2 FUSION ASSAYS ................................................................................................ 43

2.2.1 In vivo mitochondrial fusion assay ............................................................. 43

2.2.2 In vitro liposome fusion assay .................................................................... 43

2.3 MITOCHONDRIAL ASSAYS ................................................................................. 44

viii

2.3.1 Mitochondrial purification .......................................................................... 44

2.3.2 Oxidative phosphorylation ......................................................................... 45

2.3.3 Mitochondrial ATP measurements ............................................................. 45

2.4 CYCLOHEXIMIDE CHASE ............................................................................... 45

2.5 AUTOPHAGY ASSAYS ..................................................................................... 46

2.5.1 Determination of cell viability ................................................................... 46

2.5.2 Atg8/Atg8-PE analysis ............................................................................... 46

2.5.3 Vacuolar internalization of FM 4-64 .......................................................... 46

CHAPTER 3 PHOSPHATIDYLSERINE DECARBOXYLASE 1 (PSD1)

PROMOTES MITOCHONDRIAL FUSION BY REGULATING THE

BIOPHYSICAL PROPERTIES OF THE MITOCHONDRIAL MEMBRANE

AND ALTERNATIVE TOPOGENESIS OF MITOCHONDRIAL GENOME

MAINTENANCE 1 (MGM1) .................................................................................... 47

3.1 ABSTRACT .......................................................................................................... 47

3.2 INTRODUCTION ................................................................................................. 48

3.3 RESULTS ............................................................................................................... 50

3.3.1 Psd1 is required for normal mitochondrial morphology ............................ 50

3.3.2 Psd1 is required for proper mitochondrial fusion during yeast mating ...... 52

3.3.3 Phospholipid composition affects the rate of lipid mixing ......................... 54

3.3.4 Psd1 is required for proper mitochondrial activity ..................................... 56

3.3.5 Ethanolamine cannot rescue Δpsd1 mitochondrial-specific defects .......... 58

3.3.6 s*Mgm1 can rescue mitochondrial aggregation but not the

glycerol growth defect in Δpsd1 cells ......................................................... 59

ix

3.3.7 Psd1 regulates alternative topogenesis of Mgm1 ....................................... 61

3.4 DISCUSSION ....................................................................................................... 64

CHAPTER 4 MITOCHONDRIAL PHOSPHATIDYLETHANOLAMINE

IS IMPORTANT FOR STARVATION-INDUCED AUTOPHAGY

IN YEAST ................................................................................................................... 70

4.1 ABSTRACT .......................................................................................................... 70

4.2 INTRODUCTION ................................................................................................. 71

4.3 RESULTS .............................................................................................................. 73

4.3.1 Psd1 is required for proper autophagy in yeast ........................................... 73

4.3.2 The loss of Psd1 results in a reduction of autophagic bodies

in the yeast vacuole ..................................................................................... 74

4.3.3 Ethanolamine cannot rescue the autophagic defects in Δpsd1 cells ........... 76

4.4 DISSCUSSION ..................................................................................................... 78

CHAPTER 5 CONCLUDING PERSPECTIVES .......................................................... 81

5.1 BRIEF SUMMARY OF RESULTS ...................................................................... 81

5.2 PERSPECTIVES ................................................................................................... 82

5.2.1 Phosphatidylethanolamine in mitochondrial membrane dynamics ............ 82

5.2.2 Mitochondrial lipids in mitochondrial biology ........................................... 85

5.2.3 The specific role(s) of mitochondrial phosphatidylethanolamine

in autophagy ................................................................................................ 87

5.2.4 The composition of the mitochondrial membrane in autophagy ................ 89

5.3 CONCLUSION ..................................................................................................... 92

REFERENCES .................................................................................................................. 94

x

LIST OF FIGURES

Figure 1-1: Proposed model for yeast mitochondrial fission ......................................... 5

Figure 1-2: Schematic of the yeast mitochondrial fusion machinery ........................... 9

Figure 1-3: Alternative topogenesis of Mgm1 ................................................................. 11

Figure 1-4: Schematic of Mgm1-mediated inner mitochondrial membrane

fusion .............................................................................................................. 13

Figure 1-5: Different forms of autophagy ....................................................................... 18

Figure 1-6: Induction of autophagy ................................................................................. 20

Figure 1-7: Order of protein recruitment to the phagophore assembly site

in yeast ............................................................................................................ 22

Figure 1-8: The two ubiquitin-like conjugation systems in autophagy ........................ 23

Figure 1-9: Simplified overview of phosphatidylethanolamine biosynthesis in

Saccharomyces cerevisiae .............................................................................. 31

Figure 1-10: Phosphatidylethanolamine biosynthesis by phosphatidylserine

decarboxylation and the Kennedy pathway ................................................ 32

Figure 1-11: Domain organization of phosphatidylserine decarboxylases ................... 33

Figure 1-12: Phosphatidylserine decarboxylase proenzyme maturation ...................... 34

Figure 1-13: Phosphatidylserine decarboxylase enzymology ......................................... 35

Figure 3-1: Psd1 is required for normal mitochondrial morphology ........................... 51

Figure 3-2: Psd1 is required for mitochondrial fusion during yeast mating ............... 53

Figure 3-3: Liposomes with lipid compositions similar to ΔΔpsd1 mitochondria

have a reduced rate of lipid mixing ............................................................. 55

Figure 3-4: The ΔΔpsd1 strain has defects in mitochondrial activity ............................. 57

xi

Figure 3-5: s*Mgm1 suppresses ΔΔpsd1 mitochondrial aggregation ............................. 60

Figure 3-6: Psd1 regulates Mgm1 alternative topogenesis ............................................ 63

Figure 3-7: Model of Psd1-dependent mitochondrial regulation .................................. 69

Figure 4-1: Mitochondrial phosphatidylethanolamine is important for

autophagy ....................................................................................................... 75

Figure 4-2: Reducing mitochondrial phosphatidylethanolamine reduces

autophagic bodies in the vacuole .................................................................. 77

Figure 5-1: Overexpression of Psd1 results in mitochondrial fragmentation

independent of s-Mgm1 biogenesis .............................................................. 83

xii

LIST OF ABBREVIATIONS

- N Starvation medium

5-FOA 5-Fluoroorotic acid

ATP Adenosine triphosphate

BAR Bin-Amphiphysin-Rvs

BHK Baby hamster kidney

C12E8 Octaethylene glycol monododecyl ether

CCCP Carbonylcyanide m-chlorophenylhydrazone

CDP-DAG Cytidyldiphosphate diacylglycerol

CDP-Etn Cytidyldiphosphate ethanolamine

CHO Chinese hamster ovary

CHX Cycloheximide

CL Cardiolipin

CMA Chaperone-mediated autophagy

CTP Cytidyltriphosphate

Cvt Cytoplasm-to-vacuole targeting

DAG Diacylglycerol

DRP Dynamin-related protein

EM Electron microscopy

ER Endoplasmic reticulum

ERMES Endoplasmic reticulum-mitochondria encounter structure

ETC Electron transport chain

ETM Energy transfer motif

xiii

Etn Ethanolamine

GDP Guanosine diphosphate

GED Guanosine triphosphate effector domain

GTP Guanosine triphosphate

GTPase Guanosine triphosphate hydrolase

GUV Giant unilamellar vesicle

IMM Inner mitochondrial membrane

IMS Intermembrane space

lyso-PE Lyso-phosphatidylethanolamine

MAM Mitochondrial-associated endoplasmic reticulum membrane

MD Molecular dynamics

MICOS Mitochondrial contact site

mtBFP Mitochondrial-targeted blue fluorescent protein

mtGFP Mitochondrial-targeted green fluorescent protein

MPP Mitochondrial processing peptidase

MPTP Mitochondrial permeability transition pore

mTOR Mammalian target of rapamycin

mTORC1 Mammalian target of rapamycin Complex 1

MTS Mitochondrial targeting sequence

MVB Multivesicular body

NAT Nourseothricin

OD Optical density

OMM Outer mitochondrial membrane

xiv

ORF Open reading frame

PA Phosphatidic acid

PAS Phagophore assembly site

PC Phosphatidylcholine

PE Phosphatidylethanolamine

P-Etn Phosphoethanolamine

PG Phosphatidylglycerol

PHB Prohibitin

PI Phosphatidylinositol

PI3K Phosphatidylinositol 3-kinase

PI3P Phosphatidylinositol 3-phosphate

PM Plasma membrane

pmaER Plasma membrane-associated endoplasmic reticulum

PMSF Phenylmethylsulfonyl fluoride

PS Phosphatidylserine

PSD Phosphatidylserine decarboxylase

SC Synthetic complete

SUV Small unilamellar vesicle

TM Transmembrane

TOR Target of rapamycin

TORC1 Target of rapamycin Complex 1

TPR Tetratricopeptide repeat

WT Wild type

1

CHAPTER 1

INTRODUCTION

Some material in this chapter was previously published. Springer Publishing Company, 2011,

Mitochondrial dynamics and neurodegeneration, Chapter 1, pp 1-46, The genetics of

mitochondrial fusion and fission, Eliana Y. L. Chan, Jarungjit Rujiviphat and G. Angus

McQuibban, with kind permission from Springer Science and Business Media.

1.1 Mitochondrial membrane dynamics

Mitochondria were first observed in the 1850s when cytologists discovered granular

structures in the cytoplasm of living cells. In 1857, Swiss anatomist Albert von Kölliker

described these granular structures in the sarcoplasm of insect muscle cells and showed in 1888

that these granular structures swelled in water and possessed a membrane (1). These structures

had originally been named "bioblasts" and "sarcosomes" by Richard Altmann (2) and Gustaf

Retzius (3), respectively, but in 1898, Carl Benda introduced the term "mitochondrion", derived

from the Greek words mitos meaning threaded, and chondron meaning grain (reviewed in

reference (4)). Although the term "mitochondrion" has become the most widely accepted name

for this organelle, the term "sarcosome" is still used today to describe mitochondria in muscle

cells.

2

1.1.1 The discovery of mitochondrial membrane dynamics

One of the first in-depth descriptions of mitochondria in cells was achieved by Lewis and

Lewis in 1915 (5). Using cultured chick cells, they were able to observe live and fixed

mitochondria, and could describe mitochondrial movement, quantity and dynamics (5).

However, it was not until the 1990s with the development of fluorescent probes and the

resurgence of light microscopy that the movement, fusion and fission of mitochondria in living

cells were recorded and widely accepted (6). Since the identification of mitochondrial

membrane dynamics, technological advances together with genetic, structural and biochemical

approaches have enabled the field to dissect the molecular mechanisms of mitochondrial fusion

and fission and how this delicate network is maintained. What Lewis and Lewis referred to as

mitochondrial fusion and branching/separation (5) are now known to be distinct, regulated

processes required for proper mitochondrial function and inheritance (7-10). Mitochondrial

membrane dynamics are now known to be crucial mediators of apoptosis (11,12) and play a role

in neurodegeneration (13-15).

The shape of the mitochondrial network is maintained by the regulated balance of fusion

and fission events (16-21). Accordingly, decreased fission results in elongated mitochondria

(20-24), and decreased fusion results in mitochondria that appear fragmented and/or aggregated

(16-18). Notably, the phenotype resulting from reduced mitochondrial fusion can be rescued by

reducing mitochondrial fission (17,18) and vice versa (21), strongly indicating that normal

mitochondrial morphology is maintained by the balance of fusion and fission events rather than

the absolute number of each reaction.

The pioneering discoveries of the molecular machineries of mitochondrial membrane

fusion and fission were largely derived from genetic studies in yeast, flies, worms and

3

mammalian cells (reviewed in references (25,26)). Due the simplicity of a single cell system and

powerful genetic approaches, the molecular mechanisms of mitochondrial membrane dynamics

are best characterized in yeast. The key players described in yeast mitochondrial membrane

dynamics are the dynamin-related proteins (DRPs) (26). DRPs are large guanosine triphosphate

hydrolases (GTPases) involved in several membrane remodelling events (27). They are

distinguished from other GTPases by their ability to bind to lipids, self-assemble, and to undergo

oligomerization-stimulated guanosine triphosphate (GTP) hydrolysis (27). All DRPs contain

three conserved domains: (i) the GTPase domain, (ii) the middle domain, and (iii) the GTP

effector domain (GED). The catalytic GTPase domain serves as the site for GTP binding and

hydrolysis while the middle domain and GED are essential for oligomerization and self-

assembly. The region between the middle domain and the GED serves as the site for lipid

interactions (27). The four well-characterized DRPs in yeast are vacuolar protein sorting 1

(Vps1), dynamin-related 1 (Dnm1), mitochondrial genome maintenance 1 (Mgm1) and fuzzy

onions 1 (Fzo1). Vps1 mediates Golgi vesicle formation whereas the other three DRPs function

in mitochondrial membrane dynamics. Dnm1 directly mediates the fission of mitochondria

whereas Mgm1 and Fzo1 are the key players for mitochondrial fusion (26). These DRPs are

highly conserved proteins, and their higher eukaryotic counterparts are also fission and fusion

molecules.

1.1.2 Mitochondrial membrane fission

Although the regulation of mitochondrial membrane fission differs slightly from one

organism to another, the core components required remain highly conserved. In general,

mitochondrial membrane fission requires mitochondrial fission 1 (Fis1) and the DRP

4

Dnm1/Drp1. Deleting Fis1 or Dnm1 in yeast results in elongated mitochondria (20,22,28). In

yeast, Dnm1 exists as punctate structures on the outer mitochondrial membrane (OMM) and in

the cytosol (19,29), whereas mammalian Drp1 is mostly diffuse cytoplasmic with some puncta

on mitochondria (23). Despite their differences in localization, Dnm1/Drp1 in yeast, worms and

mammals localize to the OMM at sites of mitochondrial fission (30,31), consistent with their

roles as proteins that divide mitochondria. Although homologs have been identified in many

organisms, it is the detailed biochemical and structural analyses on yeast Dnm1 and mammalian

Drp1 that have shed light on the mechanism of DRP-mediated mitochondrial membrane fission.

Like dynamins, Dnm1/Drp1 assemble into dimers/tetramers (32,33) and organize into

ring-like structures (Figure 1-1) (23,34,35). The spirals formed by Dnm1 can constrict

liposomes, reducing their diameter to that of mitochondrial constriction sites (34). Independent

studies using truncated forms of yeast Dnm1 and mammalian Drp1 identified the GED as an

important region for the formation of these higher-ordered structures (32,36). The GED of

mammalian Drp1 coordinates intra- and intermolecular interactions - it forms intermolecular

interactions with itself, and intramolecular interactions with the GTPase domain (36). It is also

sufficient for the mitochondrial targeting of Drp1 (37). One of the characteristic features of

dynamins and DRPs is oligomerization-stimulated GTP hydrolysis. In the case of Dnm1/Drp1,

self-assembly into higher order oligomers occurs after recruitment to mitochondria and is the

rate-limiting step in GTP hydrolysis (34,38).

The recruitment of Dnm1/Drp1 to mitochondria occurs via Fis1, an integral membrane

protein on the OMM (Figure 1-1) (22,35). The N-terminus of Fis1 faces the cytosol and its C-

terminus faces the intermembrane space (IMS) (22,39). Most of the protein resides in the

cytosol and is anchored to the OMM by its C-terminal transmembrane (TM) segment (40).

5

Figure 1-1: Proposed model for yeast mitochondrial fission.

In yeast, Fis1 recruits the dynamin-related protein Dnm1 to sites of mitochondrial fission via the adaptor proteins Mdv1 and Caf4. The proposed model for Dnm1-dependent mitochondrial fission is depicted.

Dnm1

Fis1

Mdv1/

Caf4

OMM

GTP binding drives

Dnm1 self-assembly

Dnm1 self-assembly

constricts the membrane

GTP hydrolysis drives

Dnm1 depolymerization

and membrane instability

Membrane fission to

relieve instability

6

Unlike Dnm1/Drp1 that appear as discrete puncta, Fis1 is uniformly distributed throughout the

mitochondrial membrane (22). It organizes into six α-helices (α1-α6) that resemble

tetratricopeptide repeat (TPR) folds (41,42). TPR motifs mediate protein-protein interactions

and are usually a part of multiprotein complexes (reviewed in reference (43)). The α2-α3 and

α4-α5 helices of Fis1/hFis1 form two TPR motifs (TPR1 and TPR2), forming a concave

hydrophobic binding pocket (42,44). Although TPR motifs are proposed to mediate protein-

protein interactions, direct binding between Fis1/hFis1 and Dnm1/Drp1 remains controversial

(45-47), and adaptor proteins are proposed to mediate binding between the two. In yeast,

mitochondrial division 1 (Mdv1) and CCR4-associated factor (Caf4) serve as adaptors to recruit

Dnm1 to mitochondria (Figure 1-1) (28,48,49), whereas in higher eukaryotes, mitochondrial

fission factor (Mff), mitochondrial dynamics proteins of 49 and 51 kDa (Mid49 and Mid51,

respectively) and ganglioside-induced differentiation-associated protein 1 (GDAP1) are the

proposed receptors for Drp1 (50-56). A proposed model for mitochondrial fission in yeast is

shown in Figure 1-1.

Despite Dnm1/Drp1 and Fis1/hFis1 being the best characterized mitochondrial fission

proteins, studies in yeast and worms indicate that they may only be responsible for OMM fission.

Mitochondria of C. elegans expressing mutant forms of Drp-1 were still connected by thin

tubules of the OMM while inner mitochondrial membrane (IMM) fission still occurred (57).

Similarly, yeast Δdnm1 and Δfis1 mutants did not achieve complete mitochondrial fission, but

had matrix separation (58). These data suggest that Dnm1/Drp1 and Fis1/hFis1 are not required

for IMM fission, and that other components mediate this process.

Yeast mitochondrial distribution and morphology 33 (Mdm33), mammalian

mitochondrial protein of 18 kDa (Mtp18) and mitochondrial targeting GxxxG motif protein

7

(MTGM) were later identified and proposed to be factors mediating IMM fission (59-62).

Mdm33 has no known mammalian homologs, whereas Mtp18 has known homologs only in

metazoans (59-61). In contrast, MTGM has homologs in eukaryotes from yeast to human.

Remarkably, all the analyzed mammalian orthologs of MTGM have 100% amino acid sequence

identity (62). Reducing Mdm33, Mtp18 and MTGM protein levels resulted in elongated

mitochondria, whereas their overexpression led to fragmentation, phenotypes resembling that of

mitochondrial fission mutants (59-62). Although the topology of Mtp18 remains unclear, it

isproposed to reside in the IMS (61). Mdm33 is an integral protein of the IMM with its C-

terminus in the IMS and its predicted N-terminal coiled-coil domains in the mitochondrial

matrix. Analysis of its oligomeric state showed that Mdm33 has homotypic interactions, likely

mediated by its coiled-coil domains. Overexpressing Mdm33 resulted in IMM fragmentation

and loss of cristae, suggesting IMM fission defects. Based on these observations, Mdm33 on

apposing IMMs are proposed to form homotypic interactions via their coiled-coil domains in the

mitochondrial matrix, thereby mediating IMM fission (59).

MTGM was identified as a human nuclear gene whose product is highly enriched in brain

tumor cell lines and tumor tissues (62). Like Mdm33, MTGM is an integral membrane protein

of the IMM. Rather than coiled-coil domains, MTGM has a highly conserved tetrad of GxxxG

motif that also mediates protein-protein interactions. MTGM-induced mitochondrial

fragmentation is Drp1-dependent, indicating that it could be a regulator of Drp1. However, its

IMM localization suggests that MTGM might mediate IMM fission (62). Additional

experiments have to be conducted to determine if MTGM is directly involved in IMM fission or

whether it is simply a regulator of Drp1. A complete review of the mitochondrial membrane

fission machinery can be found in references (25,26,63).

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1.1.3 Mitochondrial membrane fusion

Mitochondrial membrane fusion is unique and distinct from other intracellular membrane

fusion events because it requires the coordinate fusion of two separate membranes. OMM and

IMM fusions are separable events with distinct energy requirements (64), indicating the

existence of distinct OMM and IMM fusion machineries. However, they are temporally coupled,

strongly suggesting that the OMM and IMM fusion machineries must coordinate effectively to

ensure proper fusion of the organelle (26). Although some proteins of the mitochondrial fusion

machinery have been identified and characterized in higher organisms, studies in yeast have laid

the foundation for our current working model of mitochondrial membrane fusion. In yeast,

OMM fusion is mediated by Fzo1 (mitofusins 1 and 2 [Mfn1 and Mfn2, respectively] in

mammals), while IMM fusion is mediated by Mgm1 (optic atrophy 1 [OPA1] in mammals). A

fungal-specific protein, Ugo1 (Ugo is Japanese for fusion), which is a member of the

mitochondrial transporter family, serves as an adaptor protein, coordinating OMM and IMM

fusion in yeast (Figure 1-2).

1.1.3.1 Outer mitochondrial membrane fusion

Fzo1 is an OMM-localized protein with a bipartite transmembrane domain (16,65,66).

The majority of the protein and both the N- and C-terminus face the cytosol (16,65). Fzo1

contains a GTPase domain at its N-terminus (65) and a total of three heptad repeats predicted to

form coiled-coil domains that mediate stable cis and trans Fzo1-Fzo1 interactions (67). Similar

to Dnm1, self-assembly and GTPase activity are crucial for Fzo1 function (64). It is proposed

that GTP binding facilitates Fzo1 cis dimerization (68). Fzo1 dimers on opposing membranes

then associate in trans to tether the membranes and facilitate fusion (Figure 1-2) (64,68).

9

Subsequent GTP hydrolysis might induce a conformational change that results in Fzo1

ubiquitination and its proteasome-dependent degradation (68-70). Dimerization in trans appears

to be conserved, as the mammalian orthologs of Fzo1, Mfn1 and Mfn2, also oligomerize in trans

Figure 1-2: Schematic of the yeast mitochondrial fusion machinery.

The yeast mitochondrial fusion complex consists of Fzo1 and Ugo1 on the outer mitochondrial membrane (OMM) and Mgm1 on the inner mitochondrial membrane (IMM). Ugo1 serves as an adaptor, coordinating OMM and IMM fusion. The dimerization of Ugo1 and the binding of guanosine triphosphate (GTP) by Fzo1 allow Fzo1 to dimerize in cis and in trans to tether the opposing OMMs. Subsequent events leading to the fusion of the OMM are not known, but Fzo1-dependent GTP hydrolysis likely induces membrane stress and destabilization that can be relieved upon Ugo1-dependent lipid mixing of the OMM. Mgm1 also oligomerizes in cis and in trans to tether opposing IMMs. GTP hydrolysis induces a conformational change in Mgm1 that could also increase membrane stress and destabilization that can be relieved upon Ugo1-dependent IMM fusion.

OMM

IMM

Ugo1

Ugo1

Fzo1

Fzo1

s-Mgm1

l-Mgm1

10

to tether mitochondria (71-73). Moreover, in the absence of a functional GTPase domain in

Mfn1, mitochondria aggregate but maintain a uniform distance (73), suggesting that dimerization

in trans facilitates mitochondrial tethering, whereas GTP hydrolysis promotes mitochondrial

fusion.

The mechanistic details of Fzo1-mediated mitochondrial membrane fusion are poorly

described due to the lack of structural details on Fzo1. However, structural studies on the

bacterial dynamin-like protein, BDLP, which is closely related to Fzo1, might shed light on

Fzo1-mediated mitochondrial fusion. BDLP binds to and tubulates lipid bilayers and may induce

a compressed bilayer by distorting lipid tails (74,75). Guanosine diphosphate (GDP)-bound

BDLP has a lower affinity for the lipid bilayer and may possess a dimeric structure that differs

from the GTP-bound form. These data along with recent crystal structures of human dynamin 1

(76,77) suggest that GTP binding induces protein polymerization (75-77) and membrane

tubulation (75), whereas GTP hydrolysis leads to protein depolymerization (75-77), leaving a

highly curved outer leaflet in a high-energy state that can be stabilized upon fusion (75).

1.1.3.2 Inner mitochondrial membrane fusion

While OMM fusion is mediated by Fzo1, IMM fusion is dependent on Mgm1. Knocking

out Mgm1 in yeast or OPA1 in mammals results in fragmented mitochondria (78-80). Mgm1 is

a nuclear-encoded protein containing an N-terminal mitochondrial targeting sequence (MTS) that

is followed by two hydrophobic segments. It is targeted to the IMM via its MTS and undergoes

a unique form of processing known as alternative topogenesis that is dependent on adenosine

triphosphate (ATP) and a functional mitochondrial protein import machinery (81). Alternative

topogenesis of Mgm1 results in the formation of the two different isoforms of Mgm1, long-

11

Mgm1 (l-Mgm1) and short-Mgm1 (s-Mgm1) (Figure 1-3) (81). Upon import into mitochondria,

full-length Mgm1 (FL-Mgm1) can be inserted into the membrane by its first hydrophobic

segment; the subsequent cleavage of its MTS by the matrix processing peptidase (MPP) results

in the formation of l-Mgm1. In an ATP-dependent manner, FL-Mgm1 can bypass its first

hydrophobic segment and be embedded in the IMM by its second hydrophobic segment where it

can be proteolytically processed by the rhomboid intramembrane protease Rbd1/Pcp1 (82,83).

Maintaining approximately equal molar concentrations of l-Mgm1 and s-Mgm1 is crucial for

mitochondrial membrane fusion. Although both isoforms are necessary for proper fusion, a

Figure 1-3: Alternative topogenesis of Mgm1.

Mgm1 contains two N-terminal hydrophobic regions. Upon import into mitochondria, full-length Mgm1 (FL-Mgm1) can be inserted and anchored into the IMM by its first hydrophobic region. The proteolytic cleavage of its mitochondrial targeting sequence (MTS) by the mitochondrial processing peptidase (MPP) results in the formation of long-Mgm1 (l-Mgm1). Alternatively, in an adenosine triphosphate (ATP)-dependent manner, FL-Mgm1 can be inserted into the IMM by its second hydrophobic region that contains the cleavage site for the rhomboid intramembrane protease, Rbd1. Subsequent cleavage by Rbd1 results in the formation of short-Mgm1 (s-Mgm1) which is released into the intermembrane space (IMS).

IMS

matrix

FL-Mgm

1

MPP

l-Mgm

1

ATP

FL-Mgm

1

Rbd1

s-Mgm

1

IMM

12

functional GTPase domain is only necessary in s-Mgm1, suggesting that the two isoforms of

Mgm1 perform different functions (84). Additionally, while s-Mgm1 preferentially localizes to

regions where the IMM and OMM are closely apposed, l-Mgm1 is mostly localized to

mitochondrial cristae (84).

Like other DRPs, Mgm1 contains the three core dynamin domains: (i) the GTPase

domain, (ii) the middle domain and (iii) the GED. In addition, Mgm1 might also contain a lipid-

binding domain located between the middle domain and the GED (85,86). It is proposed that

Mgm1-mediated IMM fusion occurs via a two-step mechanism: IMM tethering followed by lipid

mixing (87). l-Mgm1 and s-Mgm1 associate in cis and in trans to tether opposing IMMs,

forming a tight IMM interface that promotes membrane fusion (Figure 1-4) (85-88). Subsequent

GTP binding and hydrolysis by s-Mgm1 induces a conformational change in the protein

oligomer that promotes lipid mixing (89). Interestingly, s-Mgm1 assembles into a hexameric

ring that appears to be comprised of three dimers (85,89), suggesting two levels of

oligomerization, possibly cis oligomerization on the same IMM and trans oligomerization on

opposing IMMs. It is also proposed that Mgm1, like many DRPs, might induce membrane

tubulation to increase membrane stress that can be relieved upon fusion. Although OPA1 has

been demonstrated to tubulate liposomes (90), s-Mgm1-dependent liposome tubulation occurs

only rarely (89). However, the in vitro experiment was performed in the absence of l-Mgm1

(89), and the maximum GTPase activity of s-Mgm1 is only attained in the presence of l-Mgm1

(85). Hence, further studies with both l-Mgm1 and s-Mgm1 are required to fully understand the

mechanism of Mgm1-mediated IMM fusion.

13

Figure 1-4: Schematic of Mgm1-mediated inner mitochondrial membrane fusion.

Like Fzo1, Mgm1 dimerizes in cis and in trans. l-Mgm1 and s-Mgm1 dimerize on the same membrane whereas s-Mgm1 interacts with both l-Mgm1 and s-Mgm1 on opposing membranes. Mgm1 dimerization in trans tethers the opposing IMMs. GTP binding and hydrolysis induces a conformational change in Mgm1 that is proposed to increase membrane stress and instability which can be relieved upon fusion. Although Ugo1 is required for IMM fusion, its specific role still remains to be determined.

1.1.3.3 Coordinating outer and inner mitochondrial membrane fusion

Given that mitochondria have four distinct biochemical compartments (OMM, IMS,

IMM and mitochondrial matrix), and that the disruption of this organization can lead to cell

death by apoptosis, the coordination and regulation of membrane fusion is critically important.

In yeast, a relatively well-characterized protein, Ugo1, serves a bridging function during this

complex process and is required for both OMM and IMM fusion (91). Ugo1 directly interacts

with Fzo1 and Mgm1 at its N- and C-terminus, respectively (79,92). In the absence of Ugo1, the

Fzo1-Mgm1 interaction is abolished (92). Ugo1 has three TM-spanning domains and is

classified as a mitochondrial transport/carrier protein as it contains the characteristic energy

IMM

s-Mgm1l-Mgm1

?

Ugo1Ugo1

14

transfer motifs (ETMs). Its ETMs (ETMs 1 and 2) reside in TM regions 1 and 3, and facilitate

Ugo1 dimerization, creating a 6-TM complex reminiscent of transport/carrier proteins (91,93).

Although Ugo1 is not required for OMM or IMM tethering, it is required for the lipid-mixing

step of mitochondrial membrane fusion (91).

Based on the current findings, the proposed model for mitochondrial membrane fusion

involves Fzo1, Mgm1 and Ugo1 (Figures 1-2, 1-4). Ugo1 dimerization via its ETMs likely

promotes Fzo1 cis dimerization after GTP binding (68,91,93). Since Ugo1 also physically

interacts with Mgm1 (79,92), Ugo1 dimerization likely also induces Mgm1 oligomerization in

cis, although GTP-binding might not be a pre-requisite for Mgm1 oligomerization (89). OMM

fusion proceeds when Fzo1 oligomerizes in trans to tether the opposing OMMs (64,68).

Subsequent Fzo1-dependent GTP hydrolysis likely induces a conformational change (68) that

might result in membrane stress and destabilization that can be relieved by Ugo1-dependent lipid

mixing (91). Similarly, after OMM fusion, Mgm1 likely oligomerizes in trans to tether the

opposing IMMs (85,88,89), and GTP binding and hydrolysis induces a conformational change

(89) that also likely results in membrane stress and destabilization that can be relieved upon

IMM fusion. Notably, to date, no human homolog of Ugo1 has been identified. It is proposed

that a protein with a similar function to Ugo1 is required for bridging OPA1 and Mfn1/2, and

plays a role in mammalian mitochondrial membrane fusion. Given that Ugo1 plays such a

critical function in mitochondrial membrane fusion, the identity and characterization of this key

biochemical activity is needed to fully dissect mammalian mitochondrial fusion and represents a

key missing piece of the puzzle.

15

1.1.3.4 The role of lipids in mitochondrial membrane fusion

Depending on the diameter of the hydrophilic head relative to the hydrophobic tail, lipids

can adopt different shapes, affecting the way they pack (94). This ultimately leads to different

membrane curvatures that may influence biological functions such as protein import and

membrane fusion events (94,95). There has been increasing evidence demonstrating the cross-

talk between lipid metabolism and mitochondrial membrane dynamics (96). It is proposed that

non-bilayer-forming lipids such as phosphatidylethanolamine (PE) and phosphatidic acid (PA)

generate negative membrane curvature, a characteristic that promotes membrane fusion,

including fusion of mitochondria (95). A recently identified mitochondrial-localized

phospholipase D (mitoPLD) that synthesizes PA from the hydrolysis of cardiolipin (CL) is

required for proper mitochondrial membrane fusion (97). Knocking down mitoPLD resulted in

mitochondrial fragmentation and reduced mitochondrial membrane fusion, whereas its

overexpression led to mitochondrial aggregation (97). Intriguingly, these mitochondria were

separated by a distance equivalent to half that of mitochondria tethered by heptad repeats,

suggesting that this aggregated phenotype might be a result of excess tethering but a complete

lack of lipid mixing (97). This suggests that mitoPLD might promote the lipid-mixing step of

OMM fusion.

Additionally, several recent findings suggest a functional relationship between

phospholipid metabolism, Mgm1 processing and mitochondrial morphology. In particular, CL

and PE have been implicated in proper s-Mgm1 biogenesis. Altered s-Mgm1 protein levels were

observed in cells lacking unprocessed 1 (Ups1) (98) and unprocessed 2 (Ups2) (99), protein of

relevant evolutionary and lymphoid interest (PRELI)-like proteins that regulate CL (100) and PE

(99) levels in mitochondria, respectively. The prohibitins (PHBs) Phb1 and Phb2 that form a

16

multimeric complex are proposed to provide a scaffold to increase local concentrations of PE

(101) and CL (99). PHB1 genetically interacts with UPS1 and UPS2 (99) as well as genes

directly involved in lipid biosynthesis, CL synthase, CRD1, and phosphatidylserine (PS)

decarboxylase, PSD1 (99,101), the key yeast enzyme that synthesizes PE. These results indicate

that cells cannot tolerate a simultaneous reduction in Phb1 and CL or PE, suggesting that the

function of Phb1 becomes particularly important when CL and PE levels are reduced. Both CL

(102,103) and PE (104,105) have been shown to influence the activity of the supercomplexes in

the electron transport chain (ETC) and may be required for normal mitochondrial morphology as

PHB1 also genetically interacts with several genes involved in the maintenance of mitochondrial

morphology (99). Hence, it is clear that there is still much to discover with respect to the

control, orchestration and regulation of mitochondrial membrane fusion.

1.1.4 Mitochondrial membrane dynamics and autophagy

Recently, mitochondrial membrane dynamics have been implicated in autophagy, a

quality control system (106). Autophagy, which can be induced by starvation, is a mechanism

by which the cell sequesters cytoplasmic contents into double-membrane structures known as

autophagosomes. Subsequent fusion of the autophagosomes with the vacuole (in yeast) or

lysosomes (in mammals) allows for the degradation of its contents by the vacuolar/lysosomal

hydrolases. The degraded products can then be exported to the cytosol to be reused (discussed in

more detail in section 1.2). Under conditions of nutrient deprivation, defects in autophagy result

in increased cell death (107). A recent study demonstrated that mitochondrial elongation

prevented cell death during starvation-induced autophagy (106). Upon starvation, the

recruitment of the fission protein, Drp1, to mitochondria was reduced, resulting in decreased

17

mitochondrial fission with unopposed fusion. This led to elongated mitochondria that continued

producing ATP, thereby protecting the cell from death (106). This result supports the finding

that defects in mitochondrial function impair autophagy induction and autophagic flux (108) and

are the cause of death in autophagy-deficient mutants during starvation (109). Mitochondria are

also proposed to be a source of membranes for the growing autophagosome (110,111). These

data demonstrate the relationship between mitochondrial dynamics, function and autophagy, and

suggest that mitochondrial lipids may also play critical roles in the regulation of autophagy.

1.2 Autophagy

Autophagy is a cellular quality control mechanism conserved throughout evolution. It is

required for a myriad of cellular processes including the clearance of large protein aggregates

and damaged organelles, survival during starvation and proper tissue differentiation and

development of the organism. The many biological functions of autophagy are reviewed in

references (112-114).

There are three forms of autophagy: (i) chaperone-mediated autophagy (CMA), (ii)

microautophagy and (iii) macroautophagy (Figure 1-5). In CMA, which has not been

characterized in yeast, proteins with a pentapeptide motif are recognized by the cytosolic

chaperone heat shock cognate protein of 70 kDa (Hsc70) that delivers them to lysosomes for

degradation (reviewed in reference (115)). In microautophagy, cytosolic contents are directly

engulfed by the vacuole/lysosomes by the invagination, protrusion or septation of the

vacuolar/lysosomal membrane (116,117). The third and best-studied form of autophagy is

macroautophagy (hereafter referred to as "autophagy"). During autophagy, bulk or targeted

cytosolic contents are surrounded by double-membrane structures known as phagophores which

18

mature into autophagosomes, fully enclosing their contents. The outer membrane of the

autophagosomes then fuse with the vacuole/lysosomes, allowing the inner autophagosomal

membrane and its contents to be degraded by the vacuolar/lysosomal hydrolytic enzymes. The

products, such as amino acids, can then be translocated into the cytosol to be reused by the cell

(reviewed in references (118,119)).

Figure 1-5: Different forms of autophagy.

The three forms of autophagy: (i) chaperone-mediated autophagy (CMA), (ii) microautophagy and (iii) macroautophagy. In CMA, proteins with a pentapeptide motif are recognized by the cytosolic chaperone Hsc70 that delivers them to the lysosome to be degraded. In microautophagy, cytosolic contents are directly engulfed by the vacuole/lysosomes, whereas in macroautophagy, cytosolic contents are surrounded double-membrane phagophores which mature into autophagosomes. Fusion of the autophagosomes with the vacuole/lysosomes allows for the degradation of the inner autophagosomal membrane and its contents.

vacuole/

lysosome

pentapeptide

motif

Hsc70

chaperone-

mediated

autophagy

(CMA)

microautophagy

phagophore

autophagosome

macroautophagy

autolysosome

19

1.2.1 The autophagy pathway and its molecular machinery

In the last decade, a considerable number of studies have dramatically expanded our

knowledge of the molecular machinery that orchestrates autophagy. In fact, the yeast cytoplasm-

to-vacuole targeting (Cvt) pathway, which is a biosynthetic process that delivers vacuolar

hydrolases from the cytosol to the vacuole, shares the core autophagic machinery and hence is

also considered a form of selective autophagy (120). To date, over 30 autophagy-related (Atg)

genes have been identified in yeast (107,118,121-126). The proteins of the autophagic

machinery regulate processes such as cargo recognition, phagophore expansion and even the

release of degraded products into the cytosol.

1.2.1.1 The induction of autophagy

Autophagy is activated under conditions such as nutrient starvation, when the target of

rapamycin (TOR) pathway is inactivated. In yeast, TOR inactivation results in the formation of

the Atg1 complex which consists of Atg1, Atg13 and the Atg17-Atg31-Atg29 subcomplex, and

is the trigger for the induction of autophagy (Figure 1-6) (127-131). In mammalian cells, the

induction of autophagy occurs in a slightly different manner from yeast but is also triggered by

the inactivation of the mammalian TOR (mTOR) pathway and involves the formation of the

Unc-51-like kinase (ULK) complex, the mammalian ortholog of the Atg1 complex (Figure 1-6)

(132-138). The induction of autophagy by the Atg1/ULK complex is reviewed in reference

(139).

20

Figure 1-6: Induction of autophagy.

Autophagy is induced when the target of rapamycin (TOR) pathway is inactivated. In yeast, under nutrient-rich conditions, the active TOR Complex 1 (TORC1) hyperphosphorylates Atg13 which interacts with the Atg17-Atg31-Atg29 subcomplex. TORC1-dependent phosphorylation of Atg13 prevents its association with Atg1. Upon starvation, the inactive TORC1 no longer phosphorylates Atg13, resulting in its dephosphorylation and association with Atg1. The binding between Atg13 and Atg1 enhances the kinase activity of Atg1, resulting in its autophosphorylation. The formation of the Atg1-Atg13-Atg17-Atg31-Atg29 complex initiates autophagy in yeast. In mammalian cells, ULK1/2 (orthologs of yeast Atg1) constitutively associates with the complex that consists of mammalian Atg13, FIP200 (putative homolog of yeast Atg17) and Atg101 (an Atg13 binding protein). Under nutrient-rich conditions, mammalian TORC1 (mTORC1) also associates with this complex. mTORC1 phosphorylates ULK1/2, inhibiting its kinase activity. mTORC1 also hyperphosphorylates mammalian Atg13. Upon starvation, mTORC1 dissociates from the complex resulting in the dephosphorylation of ULK1/2, thereby enhancing its kinase activity and autophosphorylation. ULK1/2 also phosphorylates mammalian Atg13 and FIP200, inducing autophagy.

nutrient-rich nutrient-deprived

TORC1Atg1Atg13

Atg31

Atg29

PPP

TORC1

Atg1P

starvation

Yeast

mTORC1FIP200

FIP200PMammals

Atg101

P

ULK1/2

Atg101

PmTORC1

starvationULK1/2

Atg13

PPP

Atg13

P

Atg17

Atg13

Atg31

Atg29

Atg17

21

1.2.1.2 Phagophore formation

After the induction of autophagy, the Atg1 complex is responsible for the recruitment of

additional Atg proteins to initiate phagophore formation at the phagophore assembly site (PAS)

(128). In yeast, the PAS is juxtaposed to the vacuole (140). The recruitment of proteins to the

PAS follows a specific order (Figure 1-7) (141) and involves the activation of a Class III

phosphatidylinositol 3-kinase (PI3K) complex that also localizes to the PAS (142) (reviewed in

reference (121)). The activity of vacuolar protein sorting 34 (Vps34), the PI3K in this complex,

results in the formation of phosphatidylinositol 3-phosphate (PI3P) that assists in the recruitment

of additional Atg proteins (143,144). In mammalian cells, the PI3K complex localizes to a

subdomain of the endoplasmic reticulum (ER) known as the omegasome that is rich in PI3P

(145,146). The omegasome is proposed to serve as a platform for the recruitment of additional

Atg proteins, the expansion of phagophores and their maturation into autophagosomes (145,147).

Although the Class III PI3K is the major contributor of PI3P during autophagy, a very recent

study demonstrated that the Class II PI3K also contributes significantly to PI3P formation during

autophagy (148).

1.2.1.3 Phagophore expansion and autophagosome formation

After phagophore formation, additional proteins orchestrate the expansion and maturation

of phagophores into autophagosomes. This process requires two ubiquitin-like conjugation

systems and results in the formation of the Atg12-Atg5/Atg16 complex (149) and the Atg8-

phosphatidylethanolamine conjugate (Atg8-PE in yeast, LC3-II in mammals) (Figure 1-8) (150).

Atg12 is first activated by the E1-like enzyme, Atg7 (151). It is then transferred to the E2-like

enzyme, Atg10, which covalently conjugates it to Atg5 (149,152). The Atg12-Atg5 conjugate

22

then forms a complex with Atg16 via the direct interaction between Atg5 and Atg16 homo-

oligomers (153,154). This large ~350 kDa multimeric complex has E3-like properties and

promotes the conjugation of Atg8/LC3 to PE (155) via the other ubiquitin-like conjugation

system in autophagy (150).

Figure 1-7: Order of protein recruitment to the phagophore assembly site in yeast.

In yeast, autophagosomes form at the phagophore assembly site (PAS) in close proximity to the vacuole. Atg17 acts as a scaffold to recruit downstream Atg proteins to the PAS. The tip of each arrow points to the protein (or lipid) that recruits, whereas the protein at the start of each arrow is the protein that is recruited. The Atg12-Atg5/Atg16 complex is likely recruited to the PAS via the direct binding of Atg5 to the phagophore membrane.

Atg1

Atg13

vacuole

phagophore assembly site (PAS)

Atg9Atg31

Atg17

Atg29

Atg14Vps15 Vps30

Vps34

PI3P PI

Atg18Atg2 Atg12Atg5Atg16

Atg3Atg8

23

Figure 1-8: The two ubiquitin-like conjugation systems in autophagy.

Two key components of autophagy are formed by ubiquitin-like conjugation systems: (i) the Atg12-Atg5/Atg16 complex and (ii) the Atg8-phosphatidylethanolamine (Atg8-PE) conjugate. Atg12 is first activated by the E1-like enzyme, Atg7. It is then transferred to the E2-like enzyme Atg10 that covalently conjugates it to Atg5, forming the Atg12-Atg5 conjugate. The Atg12-Atg5 conjugate non-covalently associates with Atg16 to form the Atg12-Atg5/Atg16 complex that acts as an E3-like enzyme for the second ubiquitin-like conjugation system. In the second system, the C-terminal arginine (Arg) residue of Atg8 is removed by the cysteine protease Atg4, exposing the penultimate glycine residue. Atg8 is then activated by Atg7, the same E1-like enzyme that also activates Atg12. Activated Atg8 is then transferred to the E2-like enzyme Atg3 and finally to the E3-like enzyme, the Atg12-Atg5/Atg16 complex, to be covalently conjugated to the lipid phosphatidylethanolamine (PE).

Atg12

Atg5

Atg16

Atg7

Atg10

Atg7

Atg3

Atg12

Atg8

Atg8 Arg

Atg4

Atg8Atg7

Atg7

Atg10Atg5Atg12 Atg10

Atg12Atg5Atg16

Atg7

Atg7

Atg3Atg8

Atg3

PE

Atg8 PE

24

The first step in the activation of Atg8 requires the cleavage of the C-terminal arginine

residue by the cysteine protease, Atg4 (Figure 1-8) (156). This exposes a glycine residue on

Atg8 that is covalently conjugated to PE through a ubiquitin-like conjugation system (156).

Upon cleavage by Atg4, Atg8 is first activated by the E1-like enzyme Atg7 which also activates

Atg12 (Figure 1-8). Activated Atg8 is then transferred to the E2-like enzyme, Atg3 (150).

Finally, the Atg12-Atg5/Atg16 complex acts as an E3-like enzyme (155), covalently conjugating

Atg8 to PE (150), allowing it to be tightly bound to the membrane (156). Notably, the removal

of PE from Atg8 is also essential for autophagy. Mutations that prevented Atg4-dependent

removal of PE from Atg8-PE resulted in impaired autophagy (156,157).

During autophagosome expansion, the Atg12-Atg5/Atg16 complex localizes to the PAS

likely through the direct binding of Atg5 to the phagophore and mediates Atg8-PE conjugation

by recruiting the E2-like enzyme Atg3 to the phagophore (158). This recruitment likely occurs

via the direct binding of Atg12 to Atg3 (Figure 1-7) (159,160). After Atg8 recruitment to the

phagophore and its conjugation to PE, Atg8/LC3 was reported to promote membrane tethering

and hemifusion (161,162), facilitating phagophore expansion (157,163). The presence of the

Atg12-Atg5/Atg16 complex and perhaps additional Atg proteins are proposed to restrict the

access of Atg4 to Atg8-PE, preventing the premature cleavage of Atg8-PE, thereby ensuring

phagophore expansion (164). Shortly before or rapidly after autophagosome formation, the Atg

proteins dissociate from the autophagosome, exposing the Atg8-PE conjugate to Atg4 (164,165).

Subsequent removal of PE from the Atg8-PE conjugate by Atg4 reverses the membrane tethering

and hemifusion activity of Atg8 (161) and releases Atg8 from the completely formed

autophagosome (157). In mammalian cells, phagophore expansion also requires the homotypic

fusion of Atg16-containing vesicles in a soluble N-ethylmaleimide sensitive fusion protein

25

attachment protein receptor (SNARE)-dependent manner (166). Collectively, these data suggest

that both the ubiquitin-like conjugation systems are directly involved in phagophore expansion

and autophagosome formation.

1.2.1.4 Autophagosome maturation and fusion with the vacuole/lysosomes

Once autophagosomes have formed, they fuse with the vacuole/lysosomes where their

contents will be degraded by the vacuolar/lysosomal hydrolases and released back into the

cytosol to be reused. As mentioned above, in yeast, autophagosome formation occurs at the

PAS, in close proximity to the vacuole (140). The fusion of autophagosomes with the vacuole is

dependent on SNARE fusion proteins such as vacuolar morphogenesis 3 (Vam3) (167,168),

Vam7 (168) and Vps10 interacting protein 1 (Vti1) (169,170). In contrast to the simple and

direct system in yeast, in mammalian cells, autophagosomes have been reported to form at

multiple sites including the ER (or omegasome) (145,146), the Golgi (171), mitochondria (111),

mitochondrial-associated ER membranes (MAMs) (172) and the plasma membrane (PM) (173).

Moreover, the maturation of mammalian autophagosomes requires the sequential fusion of

autophagosomes with endosomes (174) and multivesicular bodies (MVBs) (175) (forming

amphisomes) followed by fusion with lysosomes (forming autolysosomes) (176,177). During

this maturation process, the lumen of the amphisomes becomes acidic, and they acquire

lysosomal proteins and hydrolytic enzymes (178-180). Similar to the yeast system, the

maturation of mammalian autophagosomes also requires the concerted effort of SNARE

proteins. Specifically, the formation of amphisomes requires VAMP3 (181), whereas the

formation of autolysosomes requires VAMP7 (181,182), VAMP8 (183), and VTIB (170,183).

The formation of autolysosomes in mammalian cells is also dependent on the interaction

26

between a Tectonin-domain-containing protein, TECPR1, and the Atg12-Atg5 conjugate (184).

TECPR1 was reported to selectively bind to the Atg12-Atg5 conjugate (without Atg16) and to

PI3P, and hence is proposed to selectively tether autophagosomes and lysosomes, initiating the

formation of autolysosomes (184). Upon fusion of the autophagosomes with the

vacuole/lysosomes, the contents of the autophagosomes are degraded and exported to the

cytosol. In yeast, the export of amino acids from the vacuole to the cytosol is mediated by the

vacuolar efflux pump, Atg22 (185).

1.2.2 The lipids implicated in autophagy

As detailed above, the protein machinery of autophagy has been extensively studied, but

the role of lipids in the regulation of autophagy is just beginning to be appreciated. As indicated

previously, autophagy is activated when the TOR/mTOR pathway is inactivated (119). Hence,

the phosphoinositides involved in the TOR/mTOR pathway are critical modulators of autophagy

induction (186). Phosphoinositides also play a key role in autophagosome formation (143,145),

maturation (164) and fusion with lysosomes (184,187). Recently, PA has also been implicated in

the regulation of autophagy. Phospholipase D 1 (PLD1) that synthesizes PA from

phosphatidylcholine (PC), promotes autophagosome maturation through its direct binding with

PI3P (188). The other key lipid in autophagy is PE, and the efficiency of its conjugation to Atg8

is dependent on the composition and/or the curvature of the lipid membrane (158,189).

1.2.2.1 Phosphoinositides

Under nutrient-rich conditions, the Class I PI3K phosphorylates PI(4,5)P2, resulting in the

formation of PI(3,4,5)P3. The signalling cascade that follows activates TOR/mTOR, thereby

27

suppressing autophagy (190,191). However, phosphoinositides also play an important role

downstream of the TOR/mTOR pathway. As described above in section 1.2.1.2, PI3P plays a

central role in autophagosome formation in yeast and in mammalian cells

(143,145,146,186,192). Studies in yeast revealed that the inner autophagosomal membrane is

enriched in PI3P. Interestingly, structures that are in close proximity to the phagophore are also

enriched in PI3P, suggesting that these structures may supply membranes for phagophore

expansion (193). PI3P also promotes phagophore expansion by recruiting additional Atg

proteins (both positive and negative regulators of autophagosome formation) to the PAS and

growing phagophore (reviewed in reference (186)). Furthermore, in mammalian cells, the

localization of the PI3K complex to the omegasome during the early stages of autophagosome

formation occurs at sites rich in PI3P (145). In addition to roles in autophagosome formation

and expansion, PI3P also plays a significant role in the final step of autophagy by facilitating

autophagosome fusion with lysosomes. The mammalian protein TECPR1 is proposed to tether

lysosomes and autophagosomes by selectively binding to PI3P and the Atg12-Atg5 conjugate

(184). Furthermore, the product of PI3P phosphorylation, PI(3,5)P2, has also been implicated in

autophagosome maturation (194-196). It is becoming increasingly clear that phosphoinositides

have a variety of functions during autophagy, and phosphoinositides other than PI3P and

PI(3,5)P2 have also been implicated in autophagy (reviewed in references (186,187)).

1.2.2.2 Phosphatidic acid

Although the phosphoinositides, particularly PI3P, are the best characterized lipids

involved in autophagy, other lipids, including PA have also been implicated in the regulation of

autophagy. Firstly, PA has previously been shown to activate mTOR (197), suggesting that PA

28

likely suppresses autophagy. Accordingly, reducing PA production induces autophagy, although

this induction was independent of the mTOR pathway (198). Consistent with the role of PA in

activating mTOR, a recent study found that amino acids that activate mTOR also activate PLD1,

a PA-synthesizing enzyme (199). Upon amino acid stimulation, PLD1 re-localized to lysosomes

where it activated mTOR Complex 1 (mTORC1). This re-localization was dependent on the

kinase activity of hVps34, the PI3K associated with autophagy (section 1.2.1.2), and the PI3P

binding ability of PLD1. Abrogating the former or the latter abolished amino-acid-stimulated

mTOR activation (199). Collectively, these results indicate a role for PA in the activation of

mTOR and hence a role in the suppression of autophagy. The intracellular re-localization of

PLD1 upon stimulation and the dependence of this re-localization on the kinase activity of

hVps34 and the PI3P binding ability of PLD1 were confirmed by another study (188). However,

in this study, the cells were stimulated by starvation rather than amino acids, and PLD1 re-

localized to amphisomes instead of lysosomes, promoting autophagy (188). These results

suggest that PA promotes or suppresses autophagy depending on the stimulus. The

reproducibility of PLD1 re-localization and its dependence on hVps34 and PI3P strongly suggest

that the binding of PLD1 to PI3P regulates its function. However, further studies need to be

conducted in order to determine the specific signals that trigger PLD1 to switch from a pro- to an

anti-autophagy factor, or vice versa.

1.2.2.3 Membrane composition and/or membrane curvature

Besides the role of individual phospholipids in regulating autophagy, mounting evidence

suggests that the general composition of the membrane might also be important for autophagy.

Bax-interacting factor-1 (Bif-1) is a protein that indirectly interacts with Beclin-1, a subunit of

29

the mammalian Class III PI3K complex that promotes phagophore expansion (200). Bif-1

positively regulates the activity of the Class III PI3K and promotes autophagosome formation

upon nutrient deprivation (200). Bif-1 contains a Bin-Amphiphysin-Rvs (BAR) domain that

senses membrane curvature (201) and likely binds to negatively-charged lipids (202,203). This

suggests that the curvature and presence of negatively-charged lipids on the emerging

phagophore might be important for Bif-1-mediated autophagy. In addition, the efficiency of

Atg8 conjugation to PE is also dependent on the composition (189) and possibly the curvature

(158) of the membrane. The efficiency of Atg8 conjugation to PE increases with increasing

concentrations of PE and negatively-charged phospholipids in an in vitro reconstitution

experiment (189). Moreover, the in vivo observation that Atg8-PE conjugation is strictly

dependent on the Atg12-Atg5/Atg16 complex can only be recapitulated in vitro when the

proteins were incubated with lipid vesicles that had higher curvature (158). Since the

composition of the membrane affects membrane curvature, collectively, these data demonstrate

the importance of the lipid composition of the autophagosomal membrane in autophagy. PE is a

key lipid in autophagy and is known for its ability to form hexagonal phases that increase

membrane curvature (204). Hence, its role and function within the autophagosomal membrane

warrants further study. Further exploring the requirements and functions of the autophagosomal

membrane will shed light on the role of specific phospholipids during autophagy and is a focus

of this thesis work.

30

1.3 Phosphatidylethanolamine biosynthesis in Saccharomyces

cerevisiae

PE was first discovered by Johann Thudichum in 1884 when he successfully separated

lipid fractions from brain tissue (205). Although Thudichum named the lipid "kephalin", it is

now known that "kephalin" or "cephalin" is a mixture of phospholipids including

phosphatidylserine (PS) (206) and PE (207,208). PE was eventually purified from cephalin by

Rudy and Page in 1930 (209), and possibly even purer fractions were obtained from egg yolk by

Lea et al (210) and from bovine liver by Klenk and Dohmen (211) in 1955 (212). The structure

of PE was solved by Baer et al in 1952 (213).

In the yeast Saccharomyces cerevisiae, PE is synthesized by two major routes: (i) the

decarboxylation of PS by the IMM-localized phosphatidylserine decarboxylase 1 (Psd1) and the

Golgi/vacuole-localized Psd2, and (ii) the cytidyldiphosphate-ethanolamine (CDP-Etn) branch of

the Kennedy pathway. PE can also be synthesized via a minor route by the acylation of lyso-

phosphatidylethanolamine (lyso-PE) (Figure 1-9).

1.3.1 Phosphatidylserine decarboxylation

Like all the major phospholipids, the synthesis of PE by PS decarboxylation occurs via

the cytidyldiphosphate diacylglycerol (CDP-DAG) pathway, which begins by the synthesis of

PA. In yeast, PA is first converted to the intermediate CDP-DAG by CDP-DAG synthase, Cds1.

CDP-DAG is then converted to PS by the PS synthase choline requiring 1 (Cho1) in the ER at

specialized MAMs (214). The decarboxylation of PS to PE occurs either in the IMM by Psd1

(215) or in the Golgi/vacuole by Psd2 (216,217). PE can be methylated in three sequential steps

31

Figure 1-9: Simplified overview of phosphatidylethanolamine biosynthesis in Saccharomyces cerevisiae.

In yeast, the major routes of PE biosynthesis are the decarboxylation of phosphatidylserine (PS) by PS decarboxylases (PSDs) and by the cytidyldiphosphate-ethanolamine (CDP-Etn) branch of the Kennedy pathway. PE can also be synthesized by the acylation of lyso-phosphatidylethanolamine (lyso-PE).

by the PE methyltransferases 1 and 2 (Pem1/Cho2 and Pem2/Opi3) to form the other major

phospholipid, PC (Figure 1-10) (phospholipid biosynthesis in S. cerevisiae is reviewed in

reference (217)). The transport of PS from MAMs to mitochondria for decarboxylation by Psd1

is dependent on methionine requiring 30 (Met30), a member of the Skp-Cullin-F-box-protein

(SCF) ubiquitin ligase complex (218). In contrast, the transport of PS to the site of Psd2-

dependent PS decarboxylation is dependent on two other proteins, namely, the

phosphatidylinositol 4-kinase, staurosporine and temperature sensitive 4 (Stt4) (219), and the

phosphatidylinositol transfer/binding protein PstB2 (220). Mammalian cells lack Psd2 and hence

PS decarboxylation occurs only in mitochondria (221) and is catalyzed by mammalian PISD.

PS

lyso-PE

PSD

Etn/P-Etn

CDP-Etn (Kennedy pathway)

PE

acylation

major routes

minor route

32

Figure 1-10: Phosphatidylethanolamine biosynthesis by phosphatidylserine decarboxylation and the Kennedy pathway.

PS synthesized by the cytidyldiphosphate diacylglycerol (CDP-DAG) pathway can be decarboxylated by the IMM-localized Psd1 or by the Golgi/vacuole-localized Psd2 to form PE. PE can also be synthesized by the CDP-Etn branch of the Kennedy pathway using ethanolamine (Etn) or phosphoethanolamine (P-Etn) as substrates. The methylation of PE results in the formation another major phospholipid, phosphatidylcholine (PC). PA, phosphatidic acid; PMME, phosphatidylmonomethylethanolamine; PDME, phosphatidyldimethylethanolamine.

PS decarboxylases (PSDs) are nuclear-encoded proteins (222,223) and are synthesized as

inactive precursors. The mitochondrial-localized PSDs (type I PSDs) contain an N-terminal

MTS followed by a hydrophobic transmembrane anchor. The C-terminus of the protein contains

the characteristic LGST motif that acts as an autocatalytic cleavage site, separating the

proenzyme into the α- and β-subunit (224-226) typical of pyruvoyl-dependent decarboxylases

(Figure 1-11) (pyruvoyl-dependent decarboxylases are reviewed in reference (227)). Serinolysis

between the G and S exposes the new N-terminus of the α-subunit, an electron-withdrawing

PACds1

CDP-DAGCho1

PSPsd2

PE

PSPsd1

PE

Pem1/

Cho2

PMME

Pem2/

Opi3

PDME

Pem2/

Opi3

PC

Extra-mitochondrial

space

Golgi/

vacuole

Mitochondria

Etn P-EtnEct1

CDP-Etn

Kennedy pathway

PEEpt1Eki1

33

Figure 1-11: Domain organization of phosphatidylserine decarboxylases.

Type I PSDs contain an N-terminal MTS followed by a hydrophobic transmembrane anchor (TM). The C-terminal LGST motif is a characteristic cleavage site that separates the PSD proenzyme into the α- and β-subunit. Type II PSDs contain an N-terminal sorting signal that targets them to the endomembrane system (SS). The sorting signal is followed by a C2 domain and, although the processing of type II PSDs has not been extensively studied, they contain a C-terminal GGST motif with a similar function as the LGST motif in type I PSDs. Arrows indicate sites within the protein that undergo proteolytic processing.

pyruvoyl group (Figure 1-12). Heterodimerization between the α- and β-subunit results in the

formation of the fully functional enzyme which associates with the IMM via the β-subunit (226).

The Golgi/vacuole-localized PSDs (type II PSDs) also contain an N-terminal targeting

sequence, although this sorting signal targets them to the endomembrane system. The sorting

signal is followed by a C2 domain that usually participates in lipid and protein interactions

(Figure 1-11). The C2 domain of yeast Psd2 has been proposed to be involved in membrane

docking and/or PS transport to the site of Psd2-dependent PS decarboxylation (228). Type II

PSDs also contain a GGST motif with a similar function to the LGST motif of the type I PSDs

(228). It should be noted, however, that the specific processing of type II PSDs has not been

studied in great detail.

MTS TMType I PSDs LGST

β α

Type II PSDs GGSTSS C2

β α

34

Figure 1-12: Phosphatidylserine decarboxylase proenzyme maturation.

The steps involved in the formation of the mature α- and β-subunit. Serinolysis between the glycine and serine residues results in the α, β elimination of the β-subunit. Hydration of the hydroalanine followed by the elimination of ammonium results in the formation of the N-terminal pyruvoyl group of the α-subunit. RN, R-group on the N-terminus; RC, R-group on the C-terminus.

In S. cerevisiae, PS decarboxylation, and in particular, by Psd1, is responsible for the

majority of the PE production (229) - Psd2 activity accounts for only 4-12% of the total PSD

activity (216). Thus, mitochondria are the major source of de novo PE in yeast cells. The

catalysis of PSDs occurs in a mechanism characteristic of pyruvoyl decarboxylases. During the

formation of the enzyme-substrate complex, the α-carbon of the pyruvoyl group on the α-subunit

of PSD forms a Schiff's base with the amino group on the serine moiety of PS. Subsequent

N C CRN RC

O

N C C

O

H H2 HCH2

H

O

H

N C CRN

O

H H2

O-

mature β-subunit

+ H2O

RCC C

O

CH2

+NH3

O H

H

RCC C

O

CH3

+NH3

O H

RCC C

O

CH3

+NH4

O

mature α-subunit

glycine serine

N C CRN RC

O

C C

O

H H2

C

H

OH2

+NH3

35

decarboxylation results in the formation of an intermediate complex that is protonated at the

azomethine carbon, generating PE in a Schiff's base complex with PSD. The addition of water

across the Schiff's base regenerates the enzyme and releases the product, PE (Figure 1-13).

Figure 1-13: Phosphatidylserine decarboxylase enzymology.

The α-subunit of the PSD interacts with PS to form a Schiff's base in the enzyme-substrate complex (E-S complex). Decarboxylation followed by the protonation of the azomethine carbon results in a Schiff's base of the enzyme and product. Hydration across the Schiff's base regenerates the enzyme and releases the product, PE.

RCC C

O

CH3

O

α-subunit

Phosphatidyl-serine

Ptd

+NH3

O CH2

CH

C

O

O-

H2O

RCC C

O

CH3

Ptd

+NH

O CH2

CH

C

O

O-

E-S complex

CO2+ H+

RCC CCH3

Ptd

+NH

O CH2

CH

O-

H+

RCC CCH3

Ptd

+NH

O CH2

O

CH2

+ H2O

Ptd O CH2

CH2

+NH3

Phosphatidyl-ethanolamine

Schiff’s base

36

1.3.2 The Kennedy pathway

In addition to PS decarboxylation, PE can also be synthesized via the CDP-Etn branch of

the Kennedy pathway. Ethanolamine (Etn), which is exogenously added to the medium or

formed by the breakdown of lipids, is first phosphorylated by the Etn kinase, Eki1, forming

phosphoethanolamine (P-Etn). P-Etn can also be formed by the catabolism of sphingolipids

(230) and supplied to the Kennedy pathway (231). In what is considered the rate-limiting step

(232), P-Etn is activated by cytidyltriphosphate (CTP) to form CDP-Etn by the P-Etn

cytidyltransferase, Ect1. Finally, the Etn phosphotransferase, Ept1, catalyzes the reaction

between CDP-Etn and diacylglycerol (DAG) to form PE (Figure 1-9).

1.3.3 Alternative pathways of phosphatidylethanolamine biosynthesis

In many cell types, PS decarboxylation and the CDP-Etn branch of the Kennedy pathway

are the major sources of PE. However, PE can also be synthesized by the acylation of lyso-PE

(Figure 1-8) which in yeast, is catalyzed by the lyso-PE acyltransferases Ale1 (233,234) and

Tgl3 (235). In mammalian cells, PE can also be synthesized by base exchange, where the serine

head group in PS is exchanged for Etn. This reaction that is catalyzed by PS synthase 2 (PSS2)

is considered a relatively minor contributor of PE biosynthesis (236).

1.3.4 The different contributions of phosphatidylserine decarboxylation and

the Kennedy pathway

In E. coli, PE is synthesized exclusively by PS decarboxylation (237), whereas PE in the

trypanosome T. brucei is synthesized solely by the Kennedy pathway (238). In yeast, PS

decarboxylation is the main source of cellular and mitochondrial PE (216,229). In mammalian

37

cells, however, the relative contributions of the two pathways appear to be tissue-specific. In

many cultured cells such as Chinese hamster ovary (CHO) cells (239) and baby hamster kidney

(BHK) cells (240), PE is primarily synthesized by PS decarboxylation, whereas in rat

liver/hepatocytes (241-243) and hamster heart (244), the Kennedy pathway contributes the

majority of the total PE. While the difference in the relative contributions of the PE biosynthetic

pathways is unknown, it is speculated that the availability of the substrates serine (for PS

decarboxylation) and Etn (for the Kennedy pathway) might contribute to these observed

differences (245).

In yeast, the three different sources of PE appear to serve different purposes. Psd1-

derived PE accounts for most of the cellular and mitochondrial PE synthesized. Psd2-derived PE

is the preferred substrate for methylation into PC, and PE in the microsomal membranes is

primarily derived from the Kennedy pathway (229). Although PE synthesized by Psd2 and the

Kennedy pathway can be imported into mitochondria, import is rather inefficient, and they

contribute mostly to the OMM rather than the IMM (229). It is noteworthy that the different

pathways in mammalian cells synthesize somewhat different species of PE (245,246). PS

decarboxylation preferentially synthesizes PE with polyunsaturated acyl chains, whereas the

Kennedy pathway preferentially synthesizes PE species with mono- and di-unsaturated acyl

chains (245). Although the yeast enzymes show no difference in substrate selectivity, there is

slightly more 34:2 PE in mitochondria than in the microsomal fraction (247). Hence, it is

tempting to speculate that the different species of PE that are derived from the different pathways

could contribute to the different functions of PE in different parts of the cell.

38

1.3.5 Functions of phosphatidylethanolamine

The importance of PE is evident by the severe defects resulting from its loss or mutations

in genes affecting its metabolism. In yeast, PE makes up about 25% of the cellular lipids, and

the loss of PE synthesis results in a strict auxotrophy for Etn, a substrate of the CDP-Etn pathway

for PE synthesis (248). This suggests that a minimum amount of PE is necessary for survival.

Similarly, homozygous deletion of the Pisd gene in mice results in embryonic lethality at day 9.5

of development, demonstrating that PS decarboxylation is essential for mouse development

(249). Although the heterozygous Pisd+/- mice are viable, PE synthesis via the Kennedy

pathway is up-regulated, again demonstrating that a minimum amount of PE is necessary for

survival (249).

PE is a major constituent of biological membranes. The bacterial membrane is comprised

of ~75% PE (250), and in yeast and mammalian cells, PE is the second most abundant

phospholipid after PC, accounting for ~25% of total phospholipids (248,251,252). Due the size

of its hydrophilic head relative to its hydrophobic tail, PE is considered a non-bilayer-forming

lipid as it tends to arrange into non-lamellar hexagonal phases that induce negative membrane

curvature. Hexagonal phases are proposed to promote membrane fusion (204) and create lateral

pressure that can alter the function of membrane proteins (94). Indeed, PE-induced

conformational changes in the protein and lipid bilayer promote the fusion of post-mitotic Golgi

vesicles (253). Furthermore, reducing the amount of PE in mitochondria, the organelles that

typically contain the highest amount of PE, results in impaired mitochondrial morphology in T.

brucei (254) and in mice (249). Proper mitochondrial oxidative phosphorylation and energy

production are also dependent on PE - PE is required for the assembly and optimum activity of

the complexes in the ETC including Complex I (105,255) and Complex IV (104,105,256). PE

39

also has an unexpected role as a chaperone, assisting protein folding in E. coli (257,258). Proper

folding of the membrane protein lactose permease (LacY) is dependent on the non-bilayer-

forming property of PE (258) (reviewed in reference (259)). Recently, PE was also identified as

a factor required for the propagation and infectivity of mouse and hamster prions in the brain

(260).

In addition to the above-mentioned functions, PE also serves as a precursor for the other

major phospholipid, PC. It can be methylated in three sequential steps by methyltransferases,

resulting in the formation of PC (217). PE is also the precursor of a bacterial- and

mitochondrial-specific phospholipid, CL. Recently, a novel CL biosynthetic pathway was

discovered in bacteria (261). The CL synthase ClsC synthesizes CL using PE and

phosphatidylglycerol (PG) (261). Moreover, PE is also the source of Etn that is covalently

conjugated to various proteins as an amino acid modification. For instance, PE is the direct

precursor of Etn-phosphoglycerol that is bound to the eukaryotic elongation factor 1A (eEF1A)

(262-265). Furthermore, as noted above in section 1.2.1.3, the covalent conjugation of PE to

Atg8/LC3 is a key step in autophagy, and the disruption of this process results in impaired

autophagy (150).

1.4 Specific interests and goals

It is becoming increasingly clear that lipids are not simply membrane constituents - they

modulate the function of integral membrane proteins and play critical roles in membrane

remodelling events. In this thesis, I describe additional roles of mitochondrial-localized PE in

mitochondrial membrane dynamics and autophagy. My work demonstrates that mitochondrial

PE plays an important role in the maintenance of mitochondrial morphology by promoting

40

mitochondrial membrane fusion. Mitochondrial PE increases the biogenesis of s-Mgm1, a key

mitochondrial fusion protein and likely enhances the rate of lipid mixing during fusion. I also

identify mitochondrial PE as an important factor during autophagy in yeast. My results suggest

that mitochondrial PE might enhance autophagosome formation, expansion and/or promote the

fusion of autophagosomes with the vacuole. From my studies, I elucidate the mechanism by

which lipids might be important factors in mitochondrial membrane fusion and autophagy.

41

CHAPTER 2

MATERIALS AND METHODS

2.1 Reagents

2.1.1 Yeast strains

All strains used in these studies are derivatives of BY4741 (MATa; his3Δ1; leu2Δ0;

met15Δ0; ura3Δ0). Gene deletions and genomic tagging were performed by homologous

recombination of gene-specific PCR products. To generate the Δpsd1Δmgm1 strain harbouring

the pRS315-WT-Mgm1 or pRS315-Mgm1-G100D plasmids, a MATa Δpsd1 haploid strain was

mated with a MATα Δmgm1 haploid strain harbouring the pRS316-WT-Mgm1 plasmid. After

sporulation and tetrad dissection, ploidy and mating type were determined by a mating type test,

and the haploid Δpsd1Δmgm1 double mutant retaining the pRS316-WT-Mgm1 plasmid was

identified by viability on medium lacking uracil and resistance to the antibiotics nourseothricin

(NAT) and G418. The strain was then transformed with either the pRS315-WT-Mgm1 or the

pRS315-Mgm1-G100D plasmid. Removal of the pRS316-WT-Mgm1 plasmid was determined

by cell viability on medium lacking leucine in the presence of 5-fluoroorotic acid (5-FOA).

2.1.2 Plasmids

pYX122-mtGFP (mtGFP), pYES-mtGFP and pYES-mtBFP (galactose-inducible mtGFP

and mtBFP, respectively) were kind gifts from Dr. Benedikt Westermann and have been

previously described (266). pRS315-s*Mgm1-3xHA (s*Mgm1) was generated by cloning a

SacI/SalI double-digested fragment from pRS314-s*Mgm1-3xHA (a gift from Dr. Andreas

42

Reichert and has been previously described (84)) into pRS315. pRS315-Mgm1-G100D was also

a gift from Dr. Andreas Reichert and was previously described (81). For the overexpression of

Psd1, the open reading frame (ORF) of Psd1 was amplified using a reverse primer with a 3xHA

sequence included in the primer. The amplified ORF with a 3xHA tag was cloned into pRS415-

GPD at the PstI and XhoI restriction sites.

2.1.3 Growth conditions

Synthetic complete (or minimal) media consisted of 0.17% yeast nitrogen base, 5%

ammonium sulphate, the appropriate amino acids and 2% glucose, galactose or glycerol as the

carbon source. Rich media consisted of 1% yeast extract, 2% peptone and 2% glucose or

galactose. Ethanolamine (Etn) supplementation was added to a final concentration of 5 mM.

Starvation medium (- N) consisted of 0.17% yeast nitrogen base (without amino acid, without

ammonium sulphate) and 2% glucose as the carbon source.

2.1.4 Lipids

All lipids were purchased from Avanti polar lipids. Non-fluorescent lipids used in these

studies include 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC, 14:0 PC), 1,2-dimyristoyl-

sn-glycero-3-phosphoethanolamine (DMPE, 14:0 PE), L-α-phosphatidylinositol (liver, bovine),

1,2-dimyristoyl-snglycero-3-phospho-L-serine (DMPS, 14:0 PS), 1',3'-bis[1,2-dimyristoyl-sn-

glycero-3-phospho]-sn-glycerol (TMCL, 14:0 CL) and 1,2-dimyristoyl-sn-glycero-3-phosphate

(DMPA, 14:0 PA). Head-group-labelled fluorescent lipids include 1,2-dimyristoyl-sn-glycero-3-

phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) (14:0 NBD-PE) and 1,2-

43

dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl) (14:0

Rhodamine-PE).

2.2 Fusion assays

2.2.1 In vivo mitochondrial fusion assay

MATa cells expressing pYES-mtGFP and MATα cells expressing pYES-mtBFP were

grown to log phase in synthetic galactose medium to induce the expression of mtGFP and

mtBFP. An equal number of cells were mixed together and washed twice with synthetic glucose

medium to stop mtGFP and mtBFP production. Cells were resuspended in the same volume of

synthetic glucose medium, mixed thoroughly, and evenly spread onto a synthetic glucose agar

plate using a sterile inoculating loop. Liquid medium on the agar plate was air-dried, and the

plate was incubated for 3 h at 30°C for yeast mating. Cells were then carefully scraped off the

plate with a sterile inoculating loop, resuspended in synthetic glucose medium and analyzed by

fluorescence microscopy.

2.2.2 In vitro liposome fusion assay

Phospholipid compositions of WT and Δpsd1 mitochondria had been previously

determined (229). Briefly, liposomes with a lipid composition similar to WT mitochondria

contained (in mol%) 44% DMPC, 29% DMPE, 8% Liver PI, 4% DMPS, 6% TMCL and 9%

DMPA, whereas liposomes with a lipid composition similar to Δpsd1 mitochondria contained

52% DMPC, 9% DMPE, 19% Liver PI, 4% DMPS, 6% TMCL and 10% DMPA. For

fluorescence-labelled liposomes, 1.6% of the phosphatidylethanolamine (PE) was replaced with

0.8% NBD-PE and 0.8% Rhodamine-PE. Lipids were mixed in a glass tube and dried under a

44

gentle stream of nitrogen. Residual chloroform was removed under a rotary evaporator for 90

min. Lipid films were rehydrated to a concentration of 0.5 mM in liposome buffer (20 mM

HEPES pH 7.5, 150 mM NaCl) at 65°C for 1 h, vortexing every 15 min for homogeneity. To

make liposomes, hydrated lipids were extruded 15 times through a 1.0 µm filter membrane

(Avanti Polar Lipids). Liposome fusion was carried out in 96-well black plates in 100 µl

reactions. 1 µl of labelled liposomes was mixed with 9 µl of unlabelled liposomes in 80 µl of

liposome buffer. After reading basal fluorescence for 5 min, 10 µl of 1.0 M CaCl2 were added to

induce liposome fusion. Liposome buffer was used as a negative control, and maximum lipid

mixing was determined by adding 10 µl of 10% octaethylene glycol monododecyl ether (C12E8).

Readings were taken every minute for 90 min, and fluorescence was normalized to the initial

fluorescence reading and presented as a percentage of maximum lipid mixing. Relative rates of

lipid mixing were obtained by determining the slopes of the linear portion of the curves

immediately after the addition of CaCl2 and C12E8.

2.3 Mitochondrial assays

2.3.1 Mitochondrial purification

Cells were grown to log phase in 50 ml of rich galactose medium, and mitochondria were

purified as described in Gregg, et al with slight modifications (267). Briefly, after mitochondrial

enrichment, pellets were resuspended in 1 ml of resuspension buffer (homogenization buffer

without BSA) and gently homogenized 3 times with a tight dounce. Suspensions were then

pelleted at 3,000 x g for 5 min at 4°C. The resulting supernatants were then centrifuged at

12,000 x g for 15 min at 4°C to pellet mitochondria. Mitochondrial purity was assessed by

western blotting.

45

2.3.2 Oxidative phosphorylation

Cells were grown to log phase in rich galactose medium. 3 million cells (3 optical

densities [ODs]) were harvested per reading, pelleted and resuspended in 1 ml of fresh fully

oxygenated medium for oxygen consumption measurements using a Clark-type oxygen electrode

(Strathkelvin Instruments). 100% ethanol was added to a final concentration of 1% (v/v). After

the rate of oxygen consumption stabilized, 2 mM carbonylcyanide m-chlorophenylhydrazone

(CCCP) was added to a final concentration of 8 µM to induce maximum oxygen consumption.

2.3.3 Mitochondrial ATP measurements

Purified mitochondria were resuspended in 15 µl of 5.5% TCA, 2 mM EDTA and

incubated on ice for 10 min to extract ATP and precipitate mitochondrial proteins. Extracted

ATP was separated from precipitated proteins by centrifugation at 16,100 x g for 10 min at 4°C.

Supernatants containing ATP were collected for ATP measurements using an ATP

Determination Kit (Invitrogen) according to the manufacturer's instructions. Precipitated

proteins were washed twice with ice-cold acetone and heated at 95°C for 5 min to fully evaporate

the acetone. Pellets were then resuspended in 15 µl of reducing sample buffer and boiled for 5

min at 95°C. Protein concentration was determined using the RC DC protein assay (Bio-Rad).

2.4 Cycloheximide chase

Yeast cells were grown to log phase in rich galactose medium. Cycloheximide was

added to a final concentration of 100 µg/ml. Cells were collected at the indicated time points and

lysed by alkaline lysis (268) for western blot analysis. 0.5 ODs of cells were loaded per sample.

Mgm1 was detected using a rabbit polyclonal antibody against Mgm1, a kind gift from Dr.

46

Andreas Reichert. Fzo1 and Tom40 were detected using rabbit anti-sera against Fzo1 and

Tom40, generous gifts from Dr. Jodi Nunnari and Dr. Thomas Langer, respectively.

2.5 Autophagy assays

2.5.1 Determination of cell viability

Yeast cells were grown to log phase in synthetic minimal medium with glucose as the

carbon source. They were then transferred to - N medium, and cell viability was monitored by

the addition of an equal volume of Trypan blue. Cells positive for Trypan blue were counted as

dead cells.

2.5.2 Atg8/Atg8-PE analysis

Cells were grown in synthetic minimal medium with glucose as the carbon source,

washed twice with water and resuspended in the same amount of - N medium. At the indicated

time points, cells were harvested and lysed by alkaline lysis (268) for western blot analysis. 1

OD of cells was loaded per sample for each time point on a standard SDS-PAGE gel containing

13.5% acrylamide and 6 M urea in the separating gel (156). Atg8 and Atg8-PE were detected

using rabbit anti-serum against Atg8, a kind gift from Dr. Daniel Klionsky.

2.5.3 Vacuolar internalization of FM 4-64

Cells were grown to log phase in synthetic minimal medium with glucose as the carbon

source and the assay was performed as described in Journo, et al with the exception that FM 4-64

was added to a final concentration of 16 µM (269).

47

CHAPTER 3

PHOSPHATIDYLSERINE DECARBOXYLASE 1 (PSD1)

PROMOTES MITOCHONDRIAL FUSION BY REGULATING

THE BIOPHYSICAL PROPERTIES OF THE

MITOCHONDRIAL MEMBRANE AND ALTERNATIVE

TOPOGENESIS OF MITOCHONDRIAL GENOME

MAINTENANCE 1 (MGM1)

This research was originally published in The Journal of Biological Chemistry. Eliana Y. L.

Chan and G. Angus McQuibban. Phosphatidylserine decarboxylase 1 (Psd1) promotes

mitochondrial fusion by regulating the biophysical properties of the mitochondrial membrane

and alternative topogenesis of mitochondrial genome maintenance 1 (Mgm1). The Journal of

Biological Chemistry. 2013; 287:40131-40139. © The American Society for Biochemistry and

Molecular Biology.

3.1 Abstract

Non-bilayer-forming lipids such as cardiolipin (CL), phosphatidic acid (PA) and

phosphatidylethanolamine (PE) are proposed to generate negative membrane curvature,

promoting membrane fusion. However, the mechanism by which lipids regulate mitochondrial

fusion remains poorly understood. Here, I show that mitochondrial-localized phosphatidylserine

decarboxylase 1 (Psd1), the key yeast enzyme that synthesizes PE, is required for proper

48

mitochondrial morphology and fusion. Yeast cells lacking Psd1 exhibit fragmented and

aggregated mitochondria with impaired mitochondrial fusion during mating. I also demonstrate

that a reduction in PE reduces the rate of lipid mixing during fusion of liposomes with lipid

compositions reflecting the mitochondrial membrane. This suggests that the mitochondrial

fusion defect in the Δpsd1 strain could be due to the altered biophysical properties of the

mitochondrial membrane, resulting in reduced fusion kinetics. The Δpsd1 strain also has

impaired mitochondrial activity such as oxidative phosphorylation and reduced mitochondrial

adenosine triphosphate (ATP) levels which are due to a reduction in mitochondrial PE. The loss

of Psd1 also impairs the biogenesis of the short isoform of mitochondrial genome maintenance 1

(s-Mgm1), a protein essential for mitochondrial fusion, further exacerbating the mitochondrial

fusion defect of the Δpsd1 strain. Increasing s-Mgm1 levels in Δpsd1 cells markedly reduced

mitochondrial aggregation. My results demonstrate that mitochondrial PE regulates

mitochondrial fusion by regulating the biophysical properties of the mitochondrial membrane

and by enhancing the biogenesis of s-Mgm1. While several proteins are required to orchestrate

the intricate process of membrane fusion, I propose that specific phospholipids of the

mitochondrial membrane promote fusion by enhancing lipid mixing kinetics and by regulating

the action of pro-fusion proteins.

3.2 Introduction

Mitochondria are highly dynamic organelles, constantly undergoing fusion and fission

reactions to maintain a tubular network. While the protein machineries of mitochondrial fusion

and fission have been extensively studied, the role of lipids in regulating these processes is just

beginning to be uncovered. Non-bilayer-forming lipids such as PA, CL and PE are required for

49

proper mitochondrial morphology (97,100,249,270-272). Recently, a study examining the loss

of both CL and PE by deleting CL synthase, CRD1, and the mitochondrial PSD1, respectively,

demonstrated that both lipids are required for mitochondrial morphology and fusion (272). In a

mouse model, disrupting the mammalian homolog of PSD1, Pisd, resulted in abnormal

mitochondrial morphology (249). Furthermore, a reduction in the levels of a mitochondrial-

localized phospholipase D (mitoPLD that synthesizes PA from the hydrolysis of CL) resulted in

mitochondrial fragmentation (97). It is proposed that these non-bilayer-forming lipids induce

hexagonal phases, generating negative membrane curvature that promotes membrane fusion

(94,273). However, the mechanisms by which these lipids regulate mitochondrial dynamics

remain poorly understood.

The protein machinery of mitochondrial fusion is highly conserved from yeast to

mammals, and consists of large dynamin-like guanosine triphosphate hydrolases (GTPases). In

yeast, the outer mitochondrial membrane (OMM)-localized fuzzy onions 1 (Fzo1, mitofusins 1

and 2 [Mfn 1 and 2] in mammals) mediates OMM fusion, while the inner mitochondrial

membrane (IMM)-localized Mgm1 (optic atrophy [OPA1] in mammals) mediates IMM fusion

(65,80,274,275). A fungal-specific protein, Ugo1 (Ugo is Japanese for fusion), serves as an

adaptor, tethering Fzo1 and Mgm1, coordinating OMM and IMM fusion (92). Mgm1 exists as

two isoforms, long (l-Mgm1) and s-Mgm1 (276). A proper balance of l- and s-Mgm1 protein

levels is crucial for mitochondrial fusion - perturbing this ratio results in impaired mitochondrial

fusion (84). The formation of s-Mgm1 from full-length Mgm1 (FL-Mgm1) requires the

enzymatic activity of the mitochondrial rhomboid Rbd1/Pcp1, and is an ATP-dependent process

(81,82,276). Here, I show that Psd1 is required for maintaining mitochondrial ATP levels and

for the biogenesis of s-Mgm1 but does not impinge on Rbd1 activity. I also provide evidence

50

that Psd1 regulates mitochondrial fusion by regulating the biophysical properties of the

mitochondrial membrane, likely enhancing the rate of lipid mixing during fusion. Together, my

findings reveal the mechanisms by which Psd1-synthesized PE promotes mitochondrial fusion,

and demonstrate a complex interaction between lipid homeostasis, mitochondrial dynamics and

mitochondrial activity.

3.3 Results

3.3.1 Psd1 is required for normal mitochondrial morphology

To determine whether phospholipid composition plays a role in regulating mitochondrial

membrane dynamics, I examined the mitochondrial morphology of the Δpsd1 yeast strain. A

previous study demonstrated that the loss of Psd1 results in an alteration in the mitochondrial

phospholipid composition when cells are cultured in rich medium with lactate as a carbon

source. Despite the altered mitochondrial lipid composition, it was reported that the

mitochondrial morphology of these cells was unaffected (248). Indeed, I found that Δpsd1 cells

cultured in rich medium had mitochondria that were tubular, similar to that of wild type (WT)

cells (Figures 3-1A, B). However, upon further characterization, I found that 50% of Δpsd1 cells

had an intermediate mitochondrial morphology consisting of tubules, but lacking an organized

interconnected network that was observed in WT yeast cells (Figures 3-1A, B). Since rich

medium contains additional nutrients that can affect overall lipid metabolism, I cultured cells in

minimal medium, supplying only the nutrients necessary for survival. In contrast to cells

cultured in rich medium, 85% of Δpsd1 cells cultured in minimal medium had mitochondria that

were fragmented and aggregated (Figures 3-1A, B). This result is consistent with recent findings

that Psd1 is required for normal mitochondrial morphology (271,272). To further define the

51

C

n

v

WT

n

v

Δpsd1

A

Tubular

Intermediate

Fragmented

Aggregated

B

WT Δpsd1 WT Δpsd1 Δpsd1 + Etn

Rich Minimal

52

mitochondrial aggregation phenotype, I performed electron microscopic analysis. Unlike WT

mitochondria that were evenly distributed throughout the cytoplasm, Δpsd1 mitochondria were

clustered but not fused (Figure 3-1C). Furthermore, the IMM of Δpsd1 mitochondria was less

electron-dense, suggesting possible IMM defects (Figure 3-1C). Together, these data indicate

that Psd1 is required for normal mitochondrial morphology, highlighting the importance of

mitochondrial lipid metabolism in maintaining organellar morphology. The clustered

mitochondrial phenotype of the Δpsd1 strain resembles that of mitochondrial fusion mutants

(10), suggesting that Psd1 might regulate mitochondrial fusion.

3.3.2 Psd1 is required for proper mitochondrial fusion during yeast mating

To assess whether the mitochondrial morphology defect in Δpsd1 cells is due to a defect

in mitochondrial fusion, I performed an in vivo mitochondrial fusion assay. During yeast mating,

mitochondria fuse, and mitochondrial content mixing of the mating MATa and MATα haploids

can be used as an assay to monitor mitochondrial fusion. As a positive control, WT cells had

complete mixing of mitochondrial content (Figures 3-2A, B), indicating complete mitochondrial

fusion in this assay. In contrast, only 50% of Δpsd1 cells showed complete mitochondrial

______________________________________________________________________________

Figure 3-1: Psd1 is required for normal mitochondrial morphology.

Wild type (WT) and Δpsd1 cells were transformed with a mitochondrial-targeted green fluorescent protein (GFP) construct (mtGFP) and cultured in rich and minimal media with galactose as the carbon source. (A) Representative images of cells with tubular, intermediate, fragmented and aggregated mitochondria. (B) Quantification of the mitochondrial morphology of cells cultured in rich and minimal media. More than 300 cells were analyzed per strain. Error bars represent the standard deviation of 3 independent experiments. (C) Electron microscopic analysis of the WT and Δpsd1 strains cultured in minimal medium. Mitochondria (arrows), nuclei (n) and vacuoles (v) are indicated. Scale bars represent 500 nm. Etn, ethanolamine.

53

Figure 3-2: Psd1 is required for mitochondrial fusion during yeast mating.

(A) Representative images of cells that have undergone complete, partial and no mitochondrial fusion during yeast mating. Haploid WT and Δpsd1 strains were transformed with a galactose-inducible mtGFP or mtBFP. Strains were cultured and mated as described in section 2.2.1 and analyzed for mitochondrial content mixing. The extent of mitochondrial fusion was indicated by the amount of mixing between mtGFP and mtBFP. (B) Quantification of zygotes that have undergone complete, partial or no mitochondrial fusion. 20 zygotes were analyzed per strain.

A

Completefusion

Partialfusion

No fusion

mtGFP mtBFP Merge DIC

B

WT Δpsd1 Δpsd1 + Etn

54

content mixing while the remaining 50% had only partial mixing (Figures 3-2A, B). This result

is surprising considering the mitochondrial fragmentation and aggregation in the Δpsd1 strain.

Nevertheless, the ability of Δpsd1 mitochondria to undergo fusion indicates that Psd1 is not an

obligate member of the mitochondrial fusion machinery. However, the partial fusion phenotype

strongly implicates Psd1 as an important regulator, and that the altered phospholipid composition

of the mitochondrial membrane affects in vivo mitochondrial fusion during mating.

3.3.3 Phospholipid composition affects the rate of lipid mixing

Impaired mitochondrial fusion in the Δpsd1 strain during yeast mating suggests that the

phospholipid composition of the mitochondrial membrane plays an important role in promoting

mitochondrial fusion. To assess whether the altered lipid composition affects the biophysical

properties of the mitochondrial membrane that can influence mitochondrial fusion, I performed

an in vitro liposome fusion assay using liposomes with phospholipid compositions similar to that

of WT and Δpsd1 mitochondria (WT and Δpsd1 liposomes, respectively) (229). Upon the

addition of CaCl2 to induce fusion, Δpsd1 liposomes fused to the same extent as WT liposomes.

This indicates that altering the phospholipid composition does not affect the extent to which

liposomes fused (Figure 3-3A), and is consistent with our in vivo mitochondrial fusion assay that

Δpsd1 mitochondria can undergo complete fusion (Figure 3-2B). Interestingly, the slope of the

curve during mixing of Δpsd1 liposomes is different from that of WT liposomes (Figure 3-3A).

Since the slopes of the curves represent the relative rates of reaction, I analyzed the slopes to

determine if reduced fusion kinetics could contribute to the mitochondrial fusion defect in the

Δpsd1 strain. My analysis indicates that Δpsd1 liposomes had an ~75% reduction in their rates

of lipid mixing compared to WT liposomes (Figure 3-3B). Due to the absence of proteins in this

55

Figure 3-3: Liposomes with lipid compositions similar to ΔΔpsd1 mitochondria have a reduced rate of lipid mixing.

(A) In vitro liposome fusion was performed as described in section 2.2.2. Liposomes labelled with 7-nitro-2-1,3-benzoxadiazol-4-yl (NBD) and lissamine rhodamine B sulfonyl (Rhodamine) were mixed with unlabelled liposomes, and basal NBD fluorescence was monitored for 5 min. CaCl2 and C12E8 were added to induce fusion and maximum lipid mixing, respectively. The increase in NBD fluorescence indicates relief of NDB quenching by Rhodamine. (B) Quantification of the relative rates of lipid mixing indicated by the slopes of the curves in (A). Values represent the mean and standard deviation of three independent experiments. * p < 0.0001, N.S. Not significant (p > 0.05).

A

B

N.S.

+ CaCl2 + C12E8

60

55

50

10

5

0

*

N.S.

56

assay, my result indicates that changes in the phospholipid content of Δpsd1 mitochondria likely

alter the biophysical properties of the mitochondrial membrane, reducing the kinetics of lipid

mixing during mitochondrial fusion.

3.3.4 Psd1 is required for proper mitochondrial activity

To better define the importance of Psd1 in mitochondrial biology, I determined if Psd1 is

required for normal mitochondrial homeostatic functions such as cell growth and mitochondrial

bioenergetics. I analyzed cell growth under respiratory conditions, overall rate of oxidative

phosphorylation and mitochondrial ATP levels. Serial dilutions indicate that the Δpsd1 strain

grew slower than the WT strain on glycerol, a non-fermentable carbon source that necessitates

respiration (Figure 3-4A). To more quantitatively determine the mitochondrial metabolic defect

in the Δpsd1 strain, I analyzed oxidative phosphorylation by examining the rate of oxygen

consumed during ethanol metabolism. My data revealed that the basal rate of oxidative

phosphorylation in the Δpsd1 strain was 50% that of the WT strain (Figure 3-4B). The

mitochondrial uncoupler, carbonylcyanide m-chlorophenylhydrazone (CCCP) was added to

determine the maximum activity of the electron transport chain (ETC) (277). The maximum rate

of oxidative phosphorylation in the Δpsd1 strain was also 50% that of the WT strain (Figure 3-

4B). To determine the effect of impaired oxidative phosphorylation on energy production, a key

function of mitochondria, I analyzed the amount of ATP in purified mitochondria. Δpsd1

mitochondria had ~75% of WT mitochondrial ATP levels, indicating that Psd1 is required for the

maintenance of mitochondrial ATP (Figure 3-4C). Together, these results highlight the

importance of Psd1, and likely the phospholipid PE, in regulating crucial mitochondrial

components such as the ETC to maintain cellular respiration and energy production.

57

Figure 3-4: The ΔΔpsd1 strain has defects in mitochondrial activity.

(A) Serial dilutions of WT and Δpsd1 strains on glucose, glycerol and glycerol with Etn. (B) The rate of oxidative phosphorylation indicated by the rate of oxygen consumed (natom/minute/million cells) was measured as described in section 2.3.2. (C) Mitochondria were purified as described in section 2.3.1, and the amount of ATP (pmol/mg of protein) was measured using a luciferase assay as described in section 2.3.3. Δmgm1 is a respiratory-incompetent strain used as a negative control. (D) Western blot analysis of l-Mgm1 and s-Mgm1 protein levels in WT, Δpsd1 and Δpsd1 cell grown in the presence of Etn. (E) Quantification of western blots described in (D). All values represent the mean and standard deviation of three independent experiments. * p < 0.007, ** p < 0.02, N.S. Not significant (p > 0.05).

AGlycerol

WT

Δpsd1

Glucose Glycerol + Etn

B

*

*

**

*

N.S.

N.S.

C

*N.S.

*

WT Δpsd1 Δpsd1 + Etn

Δmgm1

* *

N.S.

WT Δpsd1 Δpsd1 + Etn

E

DWT

Δpsd1Δpsd1

+ Etn

48 kDTom40

s-Mgm1100 kD

75 kDl-Mgm1

58

3.3.5 Ethanolamine cannot rescue ΔΔpsd1 mitochondrial-specific defects

In Saccharomyces cerevisiae, the major route of PE synthesis is by the decarboxylation

of phosphatidylserine (PS) by the mitochondrial-localized Psd1 and the secretory pathway-

localized Psd2. Exogenously added ethanolamine (Etn) or Etn formed endogenously by lipid

metabolism can also be used to synthesize PE via the Kennedy pathway (Figure 1-9) (278).

Since Psd1-dependent PS decarboxylation is the major route of PE synthesis, the loss of Psd1

results in significantly reduced mitochondrial and cellular PE. It was also reported that

supplementing the Δpsd1 strain with exogenous Etn does not significantly increase

mitochondrial PE content, indicating that exogenous PE is not efficiently transported to

mitochondria (248). To directly demonstrate that the mitochondrial defects are a result of a

reduction in mitochondrial PE, I cultured Δpsd1 cells in media supplemented with Etn which

should increase extra-mitochondrial PE via the Kennedy pathway. The Δpsd1 glycerol growth

defect is partially rescued in the presence of Etn (Figure 3-4A). Interestingly, this rescue is

independent of mitochondrial bioenergetics. The addition of Etn could not rescue the Δpsd1

oxidative phosphorylation or mitochondrial ATP defects (Figures 3-4B, C). These results

strongly indicate that total cellular PE is crucial for growth, and that mitochondrial-localized PE

is required for efficient mitochondrial bioenergetics.

To assess if mitochondrial PE is important for the maintenance of proper mitochondrial

morphology, I analyzed the mitochondrial morphology of Δpsd1 cells grown in the presence of

Etn as previously described. Under these conditions, 50% of Δpsd1 cells had an intermediate

mitochondrial morphology. Mitochondria in these cells, while mostly tubular, were not

connected in a network like those of WT cells (Figures 3-1A, B). This resembles the

mitochondrial morphology of Δpsd1 cells cultured in rich medium, indicating that Psd1 is

59

required for proper mitochondrial morphology, especially under more stringent, nutrient-limiting

growth conditions.

Interestingly, it was previously observed that the loss of Psd1 results in altered ratios of l-

and s-Mgm1 (99). This could suggest that the mitochondrial morphology defect in Δpsd1 cells is

due to altered Mgm1 ratios. Thus, I determined whether the addition of Etn could alter Mgm1

protein ratios. Cells analyzed for their mitochondrial morphology were also harvested for

western blotting using Mgm1-specific antibodies. The ratio of l-Mgm1 to s-Mgm1 was not

altered in the presence of Etn (Figures 3-4D, E), indicating that the rescue of mitochondrial

fragmentation and aggregation in Δpsd1 is not a result of restoring Mgm1 ratios. The inability of

Etn to fully restore the tubular mitochondrial network and Mgm1 ratios suggests that the

imbalance of l- and s-Mgm1 protein levels contributes to the mitochondrial morphology defect in

the Δpsd1 strain.

3.3.6 s*Mgm1 can rescue mitochondrial aggregation but not the glycerol

growth defect in ΔΔpsd1 cells

A correct balance of l- and s-Mgm1 is required for proper mitochondrial fusion. l-Mgm1

exerts a dominant-negative effect and, increased accumulation of l-Mgm1 results in impaired

mitochondrial fusion (84). To address the possibility that the l- and s-Mgm1 protein imbalance

contributes to the mitochondrial morphology defect in the Δpsd1 strain, I artificially increased

the levels of s-Mgm1 by expressing a modified form of Mgm1 (s*Mgm1) on a plasmid (84).

Expression of s*Mgm1 in the Δpsd1 strain markedly reduced mitochondrial aggregation and

increased the number of cells with a tubular mitochondrial network (Figure 3-5A). Culturing

Δpsd1 cells expressing s*Mgm1 in Etn showed no further rescue of mitochondrial morphology

60

A

WT + vector

Δpsd1 + vector

Δpsd1 +vector+ Etn

Δpsd1 + s*Mgm1

Δpsd1 + s*Mgm1

+ Etn

100 kD75 kD48 kD

s*Mgm1

Tom40

WT + ve

ctor

Δpsd1 +

vecto

r

Δpsd1 +

s*Mgm1

Δpsd1 +

vecto

r + Etn

Δpsd1 +

s*Mgm1

+ Etn

C

B

WT + vector

Glucose

Δpsd1 + vector

WT + s*Mgm1

Δpsd1 + s*Mgm1

Glycerol

61

(Figure 3-5A). While s*Mgm1 could rescue the mitochondrial network and aggregation, serial

dilutions indicated that Δpsd1 cells expressing s*Mgm1 still exhibited impaired growth on

glycerol (Figures 3-5B, C). These results indicate that s*Mgm1 cannot restore mitochondrial

activity in Δpsd1 despite restoring mitochondrial morphology.

3.3.7 Psd1 regulates alternative topogenesis of Mgm1

My data indicate that the mitochondrial morphology defect in Δpsd1 cells is partly due to

altered l- and s-Mgm1 protein levels. To define how Psd1 regulates Mgm1, I determined

whether Psd1 regulates the biogenesis, proteolysis and/or degradation of Mgm1. Mgm1 has two

N-terminal hydrophobic regions. Insertion of the first hydrophobic region of full-length Mgm1

(FL-Mgm1) into the IMM results in the formation of l-Mgm1. In an ATP-dependent process,

FL-Mgm1 can be inserted into the membrane via the second hydrophobic region, bypassing the

first hydrophobic region (81). Subsequent proteolytic cleavage in the second hydrophobic region

of Mgm1 by the rhomboid protease Rbd1/Pcp1 results in the formation of s-Mgm1 (Figure 1-3)

(82,279). This alternative formation of l-Mgm1 and s-Mgm1 from FL-Mgm1 is known as

Mgm1 alternative topogenesis (81). It was previously shown that reducing the hydrophobicity of

the first hydrophobic region allows Mgm1 to bypass alternative topogenesis, resulting in most of

the Mgm1 protein being converted to s-Mgm1 (81). To determine if Psd1 regulates the balance

______________________________________________________________________________

Figure 3-5: s*Mgm1 suppresses ΔΔpsd1 mitochondrial aggregation.

(A) WT and Δpsd1 strains expressing mtGFP were transformed with an empty vector (vector) or s*Mgm1, and cultured in the presence or absence of Etn. Mitochondrial morphology was quantified as in Fig. 1B. Quantification represents the mean and standard deviation of three independent experiments. (B) Serial dilutions of WT and Δpsd1 cells with and without the s*Mgm1 expression plasmid grown on glucose and glycerol. (C) Western blot analysis indicating that s*Mgm1 is expressed.

62

of l- and s-Mgm1 by regulating Rbd1, I analyzed the processing of cytochrome c peroxidase 1

(Ccp1), another substrate of Rbd1. The processing of Ccp1 in the Δpsd1 strain was

indistinguishable from that of the WT strain (Figure 3-6A), indicating that Psd1 is not simply

required for the activity of Rbd1 but specifically regulates Mgm1. To determine if Psd1

regulates Mgm1 degradation, I performed a cycloheximide (CHX) chase. Consistently, I show

that the loss of Psd1 results in an increased l- to s-Mgm1 protein ratio (Figure 3-6B). To

determine if this altered ratio is due to altered Mgm1 degradation, I analyzed the change in s-

Mgm1 compared to total Mgm1 levels as a ratio during the CHX chase. A difference in the ratio

between the WT and Δpsd1 strains would indicate that Psd1 regulates the degradation of l-Mgm1

and/or s-Mgm1. Protein ratios at the different time points were normalized to the starting protein

ratio for each strain. My analysis indicated no change in the ratio of s-Mgm1 compared to total

Mgm1 levels over the course of 24 hours during the CHX chase (Figures 3-6B, C). This result

indicates that l- and s-Mgm1 are turned over at similar rates in the WT and Δpsd1 strains, and

that differential turnover of l- and s-Mgm1 is not the cause of altered l- and s-Mgm1 protein

ratios in the Δpsd1 strain. As a positive control for the CHX chase, I also analyzed the

degradation of Fzo1. In the WT strain, Fzo1 protein levels decreased over the course of the

CHX chase, indicating that Fzo1 is rapidly degraded (Figures 3-6B, C), as previously observed

(69). Interestingly, Fzo1 degradation is not impaired in the Δpsd1 strain. This is further

evidence that Psd1 specifically regulates Mgm1 protein levels rather than globally regulating the

stability of mitochondrial proteins. Since the formation of s-Mgm1 is ATP-dependent, my result

that Psd1 is required for the maintenance of mitochondrial ATP levels (Figure 3-4C) suggests

that Psd1 regulates the ATP-dependent mechanism of s-Mgm1 biogenesis. To more definitively

show that Psd1 regulates Mgm1 alternative topogenesis, I analyzed Δpsd1Δmgm1 cells express-

63

Figure 3-6: Psd1 regulates Mgm1 alternative topogenesis.

(A) Western blot analysis of WT and Δpsd1 cells expressing HA-tagged Ccp1. (B) A cycloheximide chase was performed for 24 hours. WT and Δpsd1 cells were harvested at the indicated time points and analyzed by western blotting using Mgm1-specific antibodies and Fzo1 and Tom40 anti-sera. (C) Fold changes in s-Mgm1/total Mgm1 and Fzo1 protein levels as quantified by densitometry of the western blots described in (B). Values represent the mean and standard deviation of three independent experiments. (D) Western blot analysis of Δpsd1Δmgm1 cells expressing WT or the G100D point mutant of Mgm1.

AWT Δpsd1

48 kD

35 kD

100 kD

75 kD

i-Ccp1

m-Ccp1

48 kD

l-Mgm1

s-Mgm1

Tom40

B

CHX (h)

100 kD

75 kD

48 kD

100 kD

0 3 10 24 0 3 10 24

WT Δpsd1

l-Mgm1

s-Mgm1

Fzo1

Tom40

C

D

48 kD

Tom40

WT

G100D

s-Mgm1

100 kD

75 kDl-Mgm1

64

ing WT Mgm1 or Mgm1-G100D, a previously described point mutant of Mgm1 (81). The

G100D point mutation results in reduced hydrophobicity in the first hydrophobic region of

Mgm1, allowing it to bypass alternative topogenesis. This results in most of the Mgm1-G100D

protein being converted to s-Mgm1 (81). Consistent with previous data, the Δpsd1Δmgm1 strain

expressing a plasmid-borne copy of WT Mgm1 had less s-Mgm1 compared to l-Mgm1 (Figure

3-6D). In contrast, the expression of Mgm1-G100D in Δpsd1Δmgm1 cells resulted in most of

the Mgm1 protein being converted to s-Mgm1 (Figure 3-6D). Taken together, my findings

indicate that Psd1 regulates the balance of l- and s-Mgm1 likely by regulating the ATP-

dependent mechanism of Mgm1 alternative topogenesis.

3.4 Discussion

Mitochondrial dynamics have emerged as a critical aspect of mitochondrial biology, and

they are implicated in several cellular outputs including apoptosis and autophagy. In the last

decade, the protein machineries of membrane fusion and fission have been the focus of intense

study and the integral protein units that orchestrate these membrane dynamics have been well

characterized. The next phase of discovery will be to understand the intricate mechanisms that

regulate mitochondrial dynamics and how they are integrated into cellular signalling networks.

Here, I have characterized the role of mitochondrial lipid metabolism in regulating overall

mitochondrial biology, and more specifically membrane fusion.

Previous reports have indicated that Psd1 impacts mitochondrial morphology. I have

identified the specific mitochondrial defects that arise when mitochondrial phospholipid

composition is altered. A recent study that dramatically reduced both CL and PE demonstrated

defects in mitochondrial membrane fusion and suggested overlapping roles of these

65

phospholipids in mitochondrial activity (272). I have uncovered several mechanistic details of

mitochondrial dysfunctions when only Psd1 is removed from mitochondria. Interestingly, most

of the defects are hidden when cells are grown in rich medium, likely the result of compensatory

lipid metabolic pathways that use the additional nutrients found in bacterial and yeast extracts

used in rich medium. Indeed, I have shown that mitochondria are tubular in Δpsd1 cells grown

in rich medium, but are fragmented and aggregated in minimal medium (Figure 3-1B). As

shown by rescue experiments, mitochondrial aggregation in the Δpsd1 strain grown in minimal

medium can be rescued by the addition of Etn (Figure 3-1B). This strongly indicates that

nutrients like Etn in the rich growth medium affect the mitochondrial morphology of Δpsd1 cells.

The ability of Etn to rescue mitochondrial aggregation but not the tubular mitochondrial

network in the Δpsd1 strain could be due to increased OMM PE due to lipid transfer at

mitochondrial-endoplasmic reticulum (ER) contact sites. It was recently shown that an ER-

mitochondria encounter structure (ERMES) complex consisting of maintenance of mitochondrial

morphology 1 (Mmm1), mitochondrial distribution and morphology 10 (Mdm10), Mdm12 and

Mdm34 tethers ER and mitochondria, facilitating phospholipid exchange between these

organelles (280). It has also been shown that the secretory pathway-localized Psd2 and the

Kennedy pathway contribute more PE to the OMM than to the IMM (229). This is further

supported by my data indicating that IMM functions are not restored with Etn supplementation.

Oxidative phosphorylation and mitochondrial ATP levels, which require a functional ETC in the

IMM, are not restored in the Δpsd1 strain in the presence of Etn (Figures 3-4B, C). Indeed, it has

previously been shown that PE influences the activity of enzymes in the ETC (281). This further

stresses the importance of Psd1 in supplying mitochondrial PE to maintain proper mitochondrial

bioenergetic activity. Curiously, although Etn supplementation could not rescue mitochondrial

66

bioenergetics, it could moderately suppress the glycerol growth defect in the Δpsd1 strain (Figure

3-4A). This result strongly indicates that lipid homeostasis not only affects mitochondrial

functions, but other pathways required for proper cellular growth, bringing further attention to

the importance of Psd1 in maintaining cellular and organellar lipid homeostasis.

In addition to a role in maintaining mitochondrial function, this study reveals that Psd1

maintains proper mitochondrial morphology by regulating mitochondrial fusion. My data

strongly indicate that the mitochondrial fusion defect in the Δpsd1 strain is due to both a reduced

rate of lipid mixing from the altered biophysical properties of the mitochondrial membrane and

also impaired Mgm1-driven mitochondrial fusion. Liposomes with phospholipid compositions

similar to that of Δpsd1 mitochondria fuse to the same extent as those with compositions similar

to that of WT mitochondria, however, the rate of fusion is only ~25% that of WT (Figures 3-3A,

B). Alterations in the lipid composition can affect many properties of the lipid membrane that

are believed to be important for membrane fusion. PE is known to generate negative membrane

curvature, increase lipid-packing stress and induce the formation of hexagonal phases – all of

which are properties that enhance membrane fusion (204,282,283). Here, I show that these

alterations to the biophysical properties of liposomes with a lipid composition similar to that of

the mitochondrial membrane reduce their fusion kinetics. However, this in vitro experiment was

performed in the absence of proteins and a second bilayer, factors that promote mitochondrial

fusion. Nevertheless, my result suggests that the lipid composition is unlikely to regulate the

extent of fusion, but might regulate the kinetics of fusion. To my knowledge, this is the first

indication that the kinetics of mitochondrial fusion are regulated by its phospholipid

composition.

67

The ability of s*Mgm1 to significantly reduce mitochondrial aggregation in the Δpsd1

strain strongly indicates that Psd1 regulates mitochondrial morphology in part by regulating

alternative topogenesis and the biogenesis of Mgm1. Although I expected the combination of

s*Mgm1 and Etn supplementation to fully rescue the mitochondrial morphology defect in the

Δpsd1 strain, I observed no significant improvement in the presence of Etn when Δpsd1 cells

expressed s*Mgm1 (Figure 3-5A). Although s*Mgm1 expression might have rescued the

imbalance of l-Mgm1 and s-Mgm1 in Δpsd1 cells, it is unlikely to restore the phospholipid

balance of the mitochondrial membrane. This is compelling evidence that Psd1 plays many non-

overlapping roles in mitochondrial regulation, all of which are critical in mitochondrial biology

and further substantiates the concept that the phospholipid composition of the IMM is critical for

proper mitochondrial activity.

Interestingly, I observed differing severities of the Δpsd1 mitochondrial morphology

defect. When Δpsd1 cells expressed the HIS3 gene (Figure 3-1B), the morphology defect was

more severe than when they expressed both the HIS3 and LEU2 genes (Figure 3-5A). Only 50%

of Δpsd1 cells had fragmented and aggregated mitochondria when expressing the HIS3 and

LEU2 genes compared to 85% when expressing only the HIS3 gene (Figures 3-5A, 3-1B,

respectively). This is strong indication that lipid homeostasis is closely linked to amino acid

biosynthesis. Indeed, leucine was previously identified as a precursor of phospholipids such as

PS, PE, and phosphatidylcholine (PC) (284). It is possible that the ability to synthesize leucine

allowed for the conversion of some leucine to PE, compensating for the reduction in PE levels in

the Δpsd1 strain.

This current study reveals that Psd1 regulates mitochondrial morphology by enhancing

the rate of lipid mixing during mitochondrial fusion. I also show that Psd1 plays crucial roles in

68

maintaining mitochondrial functions such as oxidative phosphorylation and maintaining ATP

levels. In addition, impaired Mgm1 alternative topogenesis in Δpsd1 cells impedes s-Mgm1

biogenesis, further exacerbating the mitochondrial fusion defect. A model of Psd1-dependent

mitochondrial regulation is shown in Figure 3-7. This work highlights the dual function of Psd1

in regulating mitochondrial fusion by enhancing lipid kinetics and the biogenesis of a

mitochondrial fusion protein.

While studies in the last decade have focused on the protein machinery of mitochondrial

dynamics, several recent reports have brought lipid homeostasis to the forefront of mitochondrial

biology. My work and work of others have identified critical roles of phospholipids in

maintaining cellular and organellar competence, highlighting their critical biological importance.

69

Figure 3-7: Model of Psd1-dependent mitochondrial regulation

PE generates negative membrane curvature, promoting fusion. In addition, the headgroup of PE provides favourable kinetic conditions. Since PE is known to regulate complexes of the electron transport chain (ETC), energy production is impaired with reduced PE, further contributing to the mitochondrial fusion defect by impairing s-Mgm1 protein biogenesis.

Non-bilayer-forming lipidse.g. PE, PA, CL

Bilayer-forming lipidse.g. PS, PI, PC

Increased curvature,favourable kinetic properties

ETCs-Mgm1

ATP

Mgm1

PS

Psd1

Endoplasmic reticulum

PE

70

CHAPTER 4

MITOCHONDRIAL PHOSPHATIDYLETHANOLAMINE IS

IMPORTANT FOR STARVATION-INDUCED AUTOPHAGY IN

YEAST

4.1 Abstract

During autophagy, contents of the cytosol are engulfed by double-membrane structures

known as autophagosomes. Subsequent fusion of the autophagosomes with the

vacuole/lysosomes enables the degradation of their contents, allowing for the recycling of

nutrients. The process of autophagy is of paramount importance when nutrients are limited - the

failure to undergo autophagy during nutrient starvation has been associated with increased cell

death. In this study, I demonstrate that mitochondrial phosphatidylethanolamine (PE) is

important for starvation-induced autophagy in yeast. Cells lacking phosphatidylserine

decarboxylase 1 (Psd1), the key yeast enzyme that synthesizes PE, die more quickly and show a

reduction in autophagy-related 8-phosphatidylethanolamine (Atg8-PE) levels compared to wild

type (WT) cells upon starvation. In addition, the Δpsd1 strain has a reduction in the number of

vacuolar autophagic bodies. Importantly, the addition of ethanolamine (Etn), which should

increase extra-mitochondrial PE, cannot rescue these defects, indicating that mitochondrial PE

plays an important role during autophagy. Given that the source of membranes for the

autophagosomes remains highly debated, my results support recent evidence that mitochondria

supply membranes to autophagosomes during starvation-induced autophagy. In addition, the

observed reduction in the number of vacuolar autophagic bodies in the Δpsd1 strain suggests that

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mitochondrial PE might be important for autophagosome formation, expansion and/or fusion

with the vacuole.

4.2 Introduction

Autophagy is a highly conserved self-degradation mechanism that serves as a means of

cellular quality control through the removal of damaged organelles and large protein aggregates.

It also serves as a means of nutrient recycling which is particularly important during times of

starvation. During autophagy, double-membrane structures known as phagophores form around

the cytosolic cargo bound for degradation. The phagophores mature into autophagosomes and

subsequently fuse with the vacuole (in yeast) or lysosomes (in mammals) where

vacuolar/lysosomal hydrolytic enzymes degrade their contents (described in detail in section 1.2,

reviewed in references (118,119)).

The complex process of autophagy is orchestrated by a group of proteins collectively

known as the autophagy-related (Atg) proteins. To date, over 30 Atg genes have been identified

in yeast (107,118,121-126). A key step in the progression of autophagy is the covalent

conjugation of the ubiquitin-like protein Atg8 to the lipid PE (150). Atg8 is first proteolytically

processed by Atg4 at the C-terminus, exposing a glycine residue that is then convalently

conjugated to PE by a ubiquitin-like conjugation system (Figure 1-8) (156). After Atg4-

dependent cleavage, Atg8 is activated by Atg7, an E1-like enzyme. It is then transferred to Atg3,

an E2-like enzyme (150). The Atg12-Atg5/Atg16 complex, itself also formed by a ubiquitin-like

conjugation system (Figure 1-8) (149), serves as an E3-like enzyme that completes the covalent

conjugation of Atg8 to PE (155). The formation of the Atg8-PE conjugate is a crucial step in

autophagy - depleting cellular PE in flies and yeast results in impaired autophagy (285,286).

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Besides PE, other lipids also play very important roles in autophagy (reviewed in

references (186,187)). The activation of autophagy is controlled by the target of rapamycin

(TOR) pathway. Activation of TOR by amino acids suppresses autophagy, whereas the

inactivation of TOR by starvation (or with the drug rapamycin) promotes autophagy. Since

phosphoinositides and their modifying enzymes play an indispensible role in the TOR pathway,

it should come as no surprise that they also play crucial roles in autophagy. The various

phosphorylated derivatives of phosphatidylinositol (PI) have been implicated both in the

activation and suppression of autophagy (reviewed in references (186,187)). In addition to PE

and the phosphoinositides, recently, phosphatidic acid (PA) has also been implicated in

autophagy. Phospholipase D 1 (PLD1), an enzyme that synthesizes PA from

phosphatidylcholine (PC), was shown to influence autophagy by promoting the biogenesis and

maturation of autophagosomes (188). However, a previous study demonstrated that PLD1

activates the mammalian TOR (mTOR) pathway in response to amino acid stimulation (199).

These data suggest that PA might function in both the activation and suppression of autophagy

depending on nutrient conditions, highlighting the importance of lipids in this pathway.

Recently, the mitochondrial-specific phospholipid, cardiolipin (CL), was also shown to

promote autophagy (287) and mitophagy (288), the specific clearance of damaged mitochondria

by autophagy. Under normal conditions, CL resides mostly on the inner mitochondrial

membrane (IMM). However, treating cells with mitochondrial stressors resulted in the

externalization of CL from the IMM to the outer mitochondrial membrane (OMM) where it

served as a signal to recruit microtubule-associated protein 1A/1B-light chain 3 (LC3), a

mammalian ortholog of yeast Atg8 to promote mitophagy (288). CL also binds to the immunity-

related guanosine triphosphate hydrolase (GTPase), IRGM, to promote autophagy. Non-CL-

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binding isoforms of IRGM could not induce autophagy (287). These results suggest that

mitochondrial lipids may play a role in the regulation of autophagy. To further investigate the

role of mitochondria and mitochondrial lipids in autophagy, I studied the phenotypes resulting

from the loss of Psd1, the IMM-localized enzyme that synthesizes PE. I demonstrate that

mitochondrial PE is an important factor during autophagy likely by enhancing autophagosome

formation, expansion and/or fusion with the vacuole in yeast. Upon starvation, cell survival and

Atg8 lipidation are impaired in cells lacking Psd1. The loss of Psd1 also results in a reduction in

the number of autophagic bodies in the vacuole, suggesting that there may be fewer

autophagosomes in Δpsd1 cells, or that the fusion of autophagosomes with the vacuole is

impaired in the Δpsd1 strain. More importantly, these defects are specifically due to a reduction

in mitochondrial PE, as the addition of Etn, which should increase extra-mitochondrial PE, could

not rescue these defects. My results support recent findings that mitochondria may supply

membranes for the growing autophagosome (110,111) and reveal that the biophysical properties

of the mitochondrial membrane may be important for the fusion of autophagosomes with the

vacuole/lysosomes during starvation-induced autophagy.

4.3 Results

4.3.1 Psd1 is required for proper autophagy in yeast

Under conditions of nutrient starvation, yeast cells defective in autophagy lose the ability

to survive (107). To determine whether mitochondrial PE is required for autophagy, yeast cells

were cultured in synthetic complete (SC) medium, transferred to nitrogen starvation medium (-

N), and cell death was monitored by Trypan blue staining as described in section 2.5.1. As

expected, the autophagy-defective Δatg5 strain had increased cell death upon nitrogen starvation

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compared to the WT strain (Figure 4-1A). Interestingly, an increase in cell death was observed

in the Δpsd1 strain, suggesting that Psd1 is required for proper autophagy (Figure 4-1A). To

more specifically determine if autophagy is impaired in the Δpsd1 strain, I analyzed the

conversion of Atg8 to Atg8-PE, the active form of Atg8 during autophagy (150). Consistent

with previous data, before starvation, little to no Atg8 was expressed (Figure 4-1B). However,

1h after nitrogen starvation, Atg8 was expressed at a much higher level in the WT, Δpsd1, and

Δatg5 strains (Figure 4-1B). Furthermore, in WT cells, 60% of Atg8 was expressed as Atg8-PE

1h after starvation and, 2h after starvation, 70% of Atg8 was expressed as Atg8-PE (Figures 4-

1B, C). In contrast, Δpsd1 cells had reduced amounts of Atg8-PE compared to WT cells; only

35% and 50% of Atg8 were expressed as Atg8-PE at the 1h and 2h time points, respectively

(Figures 4-1B, C). These data indicate that autophagy is impaired in the absence of Psd1.

4.3.2 The loss of Psd1 results in a reduction of autophagic bodies in the yeast

vacuole

During autophagy, in the presence of the protease inhibitor phenylmethylsulfonyl

fluoride (PMSF), autophagic bodies accumulate inside the vacuole of WT cells (107). To further

understand the requirement for Psd1 in autophagy, I analyzed the vacuolar internalization of the

dye FM 4-64 upon starvation. This assay takes advantage of the fact that FM 4-64 can diffuse

from the vacuolar membrane to the membranes of the autophagic bodies in the vacuolar lumen

(269). Cells were first cultured under conditions where FM 4-64 accumulates on the vacuolar

membrane. They were then transferred to - N medium in the presence of PMSF to inhibit the

degradation of autophagic bodies, and the staining of FM 4-64 was analyzed by fluorescence

microscopy. A reduction in the vacuolar internalization of FM 4-64 in the Δpsd1 strain when

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Figure 4-1: Mitochondrial phosphatidylethanolamine is important for autophagy.

Cells were cultured in synthetic complete (SC) medium and transferred to starvation (- N) medium. (A) Cell death was monitored by Trypan blue staining as described in section 2.5.1. More than 250 cells per strain were counted at each time point. (B) Western blot analysis using an anti-serum against Atg8. Starved cells were harvested at the indicated time points and lysed by alkaline lysis. (C) Quantification of three independent western blots described in (B) by densitometry. Values represent the mean and standard deviation of three independent experiments. * p < 0.02, ** p < 0.05.

A

B- N (h)

WT Δpsd1 Δatg50

Atg8Atg8-PE

Cdk

2Δpsd1 + Etn

17 kD

48 kD35 kD

0 2 0 2 0 21 1 1 1

C

**** *

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compared to the WT strain suggests a reduction in the number of autophagic bodies inside the

vacuole. 3h after starvation, 65% of WT cells had fully internalized FM 4-64 into the vacuolar

lumen, with no observable FM 4-64 staining on the vacuolar membrane. As a negative control, I

analyzed the vacuolar internalization of FM 4-64 in the autophagy-deficient Δatg8 strain. In

contrast to the WT strain, less than 10% of Δatg8 cells had complete vacuolar internalization of

FM 4-64 (Figures 4-2A, B). Instead, FM 4-64 remained on the vacuolar membrane in more than

70% of Δatg8 cells (Figures 4-2A, B), indicating impaired autophagy. Consistent with my

previous findings that Psd1 is required for proper autophagy, less than 20% of Δpsd1 cells had

complete vacuolar internalization of FM 4-64 (Figures 4-2A, B). However, 60% of the cells had

partial internalization of FM 4-64, with staining in the vacuolar lumen but also on the vacuolar

membrane, suggesting that there are fewer autophagic bodies in the vacuole of Δpsd1 cells as

compared to WT cells (Figures 4-2A, B). Interestingly, in all three assays (cell viability, Atg8-

PE/Atg8 expression and FM 4-64 vacuolar internalization), the autophagy defect in the Δpsd1

strain is less severe than the autophagy-deficient controls (Figures 4-1, 4-2). This suggests that

Psd1 is not essential for autophagy but instead enhances autophagy.

4.3.3 Ethanolamine cannot rescue the autophagic defects in ΔΔpsd1 cells

As previously described in sections 1.3.4 and 3.3.5, Psd1-dependent phosphatidylserine

(PS) decarboxylation is the major route of PE synthesis in S. cerevisiae. Hence, the loss of Psd1

results in a significant reduction in mitochondrial and cellular PE. Exogenously added Etn can

be used to synthesize PE via the Kennedy pathway (Figure 1-9), however, PE derived from the

Kennedy pathway is not efficiently transported to mitochondria (247). To determine whether the

observed autophagy defects are a result of reduced mitochondrial PE, Δpsd1 cells were cultured

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B

WTΔpsd1

- N 1h - N 3h

Δpsd1 +

EtnΔatg8 W

TΔpsd1

Δpsd1 +

EtnΔatg8

A

WT

Δpsd1

Δpsd1 + Etn

Δatg8

FM 4-64 DIC Overlay- N 3h- N 1h

DIC OverlayFM 4-64

WT

Δpsd1

Δpsd1 + Etn

Δatg8

78

in media supplemented with Etn which should have little effect on the level of mitochondrial PE,

but increase extra-mitochondrial PE via the Kennedy pathway. Surprisingly, neither the Atg8-

PE to total Atg8 ratio nor the FM 4-64 internalization defects could be rescued in the presence of

Etn (Figures 4-1B, C, 4-2). This result strongly suggests that mitochondrial PE plays an

important role in autophagy. Given that mitochondrial dynamics (106), function (109) and more

recently, the mitochondrial membrane (111) have been implicated in autophagy, my previous

findings that Psd1 influences all three aspects of mitochondrial biology (Chapter 3) imply that

Psd1 might play a critical role in autophagy.

4.4 Discussion

In recent years, the role of autophagy in cellular health and homeostasis has become the

subject of intense research. In the last decade, researchers have dissected the pathway of

autophagy, elucidating the mechanism of autophagy induction, autophagosome formation,

maturation and degradation (discussed in section 1.2 and reviewed in references (118,119,121)).

The major discoveries in the connection between lipids and autophagy involve (i) the regulation

of the TOR pathway by phosphoinositides and their modifying enzymes and (ii) the lipidation of

Atg8/LC3 by PE. Recently, PA was demonstrated to play a role of in the biogenesis and

maturation of autophagosomes, although its specific role remains unclear (188). In addition, the

______________________________________________________________________________

Figure 4-2: Reducing mitochondrial phosphatidylethanolamine reduces autophagic bodies in the vacuole

Cells were cultured and starved as described in section 4.3.2. (A) Representative images of FM 4-64 localization in the different strains at the indicated time points during starvation. (B) Quantification of cells with FM 4-64 localized in the vacuolar lumen, on the vacuolar membrane or both. More than 50 cells were analyzed per strain at each indicated time point.

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mitochondrial-enriched phospholipid, CL, has also been implicated in both autophagy (287) and

mitophagy (288). In this study, I have identified mitochondrial PE as an important factor during

autophagy in yeast.

My results reveal that the loss of Psd1, the key yeast enzyme that synthesizes PE, results

in impaired cell survival, altered Atg8-PE/Atg8 levels and a reduction in vacuolar autophagic

bodies upon starvation (Figures 4-1, 4-2). The inability of Etn to rescue the autophagic defects

in the Δpsd1 strain indicates that these defects are specifically due to a reduction in

mitochondrial PE (Figures 4-1B, C, 4-2). Although PE has previously been shown to be a

limiting factor during autophagy (285), the role of mitochondrial PE in autophagy has not been

extensively studied. Recently, mitochondrial dynamics and function have been shown to play

important roles during autophagy (discussed in section 1.1.4). Mitochondrial fusion during

starvation-induced autophagy was shown to be crucial in protecting cells from death (106).

Furthermore, mitochondrial dysfunction was shown to be the cause of death in autophagy-

deficient strains during starvation (109). My work and work of others have solidified the

requirement for Psd1 and PE in mitochondrial dynamics and function (105,249,272,289).

Hence, the observed autophagic defects resulting from the loss of Psd1 (Figures 4-1, 4-2) may be

indirect effects of impaired mitochondrial dynamics and function, and is in agreement with other

studies on the significance of mitochondrial dynamics and function in autophagy (106,109).

The observed reduction in the number of autophagic bodies in the Δpsd1 strain suggests

that autophagosome formation, expansion and/or fusion with the vacuole might be impaired. It

was recently shown that autophagosomes originate from mitochondria (111). Perhaps the

autophagic defects I observed were a result of impaired autophagosome formation from

mitochondria due to reduced mitochondrial PE in the Δpsd1 strain. In addition, in Chapter 3, I

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describe my findings that Psd1 promotes mitochondrial membrane fusion by providing

favourable lipid conditions during fusion. It is possible that Psd1 also influences autophagy by

promoting the fusion of autophagosomes with the vacuole. In support of this, a recent study

demonstrated that alterations in the membrane composition of the lysosomal/autophagic

compartments results in reduced autophagosome/lysosome fusion by up to 70% (290), indicating

that the composition of the autophagosomal membrane plays a crucial role in ensuring proper

membrane fusion during autophagy.

In sum, this work demonstrates that Psd1 is important for survival during starvation and

that mitochondrial PE plays an important role in the maintenance of Atg8-PE/Atg8 levels and the

number of vacuolar autophagic bodies during autophagy. More importantly, my results raise the

exciting possibility that autophagosome formation, expansion and/or fusion with the

vacuole/lysosomes might be influenced by the composition of the mitochondrial membrane,

possibly addressing the debate on the source of membranes for autophagosomes.

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CHAPTER 5

CONCLUDING PERSPECTIVES

5.1 Brief summary of results

Lipids are the main constituents of biological membranes and play profound roles in

biological processes including organellar dynamics and complex signalling networks. In this

thesis, I have specifically characterized the role of mitochondrial phosphatidylethanolamine (PE)

in mitochondrial fusion, mitochondrial function and autophagy. Using a series of in vitro and in

vivo studies, I demonstrate that PE plays an important role in mitochondrial membrane fusion.

The favourable biophysical properties of PE promote mitochondrial fusion likely by enhancing

the rate of lipid mixing during fusion. In addition, PE also promotes fusion by participating in

the biogenesis of the short isoform of mitochondrial genome maintenance 1 (s-Mgm1), a key

mitochondrial fusion protein. Since mitochondrial membrane fusion is proposed to maintain a

functional mitochondrial network, I also examined the role of PE in mitochondrial bioenergetics.

My findings indicate that a reduction in mitochondrial PE results in impaired oxidative

phosphorylation and reduced mitochondrial adenosine triphosphate (ATP) levels, critically

important mitochondrial homeostatic functions.

Recent studies have implicated mitochondrial dynamics and function in the regulation of

autophagy, a cellular quality control mechanism. Here, I extend these findings by demonstrating

that mitochondrial PE influences autophagy likely by promoting autophagosome formation,

expansion and/or fusion with the vacuole/lysosomes. This result highlights the intrinsic

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importance of mitochondrial PE in autophagy and implicates unique characteristics of the

mitochondrial membrane in autophagy.

5.2 Perspectives

5.2.1 Phosphatidylethanolamine in mitochondrial membrane dynamics

In recent years, the results from several studies have significantly deepened our

knowledge of the connection between mitochondrial lipid metabolism and mitochondrial

biology. Although the two non-bilayer-forming lipids PE and cardiolipin (CL) have been shown

to play important roles in mitochondrial membrane fusion (271,272), the details of how these

lipids modulate membrane dynamics have yet to be uncovered. As an extension of my studies

examining the role of PE in mitochondrial membrane fusion, I overexpressed phosphatidylserine

decarboxylase 1 (Psd1), expecting that the increase in mitochondrial PE would result in

increased mitochondrial fusion and hence an increase in highly interconnected mitochondria.

Surprisingly, I observed that the overexpression of Psd1 resulted in fragmented mitochondria

(Figure 5-1A, B), a phenotype suggesting decreased fusion or increased fission. This fragmented

phenotype is independent of s-Mgm1 biogenesis, as the expression of the two different isoforms

of Mgm1 appeared to be unaltered (Figure 5-1C). The fragmented mitochondrial phenotype is

particularly surprising as it was previously reported that the overexpression of Psd1 did not alter

the phospholipid composition of mitochondrial membranes. It was proposed that

phosphatidylserine (PS) import to mitochondria was the rate-limiting step in Psd1-dependent PE

synthesis (248). However, in that study, cells were cultured in rich medium, and their

mitochondrial morphology was not analyzed (248). Given my findings that certain

mitochondrial defects are masked when Δpsd1 cells are cultured in rich medium (section 3.3.1,

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Figure 5-1: Overexpression of Psd1 results in mitochondrial fragmentation independent of s-Mgm1 biogenesis.

Wild type (WT) cells were transformed with a mitochondrial-targeted GFP construct (mtGFP) and an empty vector or a Psd1 overexpression vector and cultured in selection medium with galactose as the carbon source. (A) Representative images of the mitochondrial morphology of cells expressing the empty vector (WT) and cells overexpressing Psd1 (Psd1 O/E). (B) Quantification of the mitochondrial morphology of cells described in (A). More than 200 cells were analyzed per strain. Error bars represent the standard deviation of three independent experiments. (C) Western blot analysis of cells expressing the empty vector (WT) or the Psd1 overexpression vector (Psd1 O/E).

Figures 3-1A, B), I believe that further exploration of the Psd1 overexpression phenotype

promises to elucidate more of the important roles that lipids play in mitochondrial membrane

dynamics. This unexpected result brings attention to the requirement for a specific amount of PE

and perhaps PS on the mitochondrial membrane. While too little PE reduced membrane

curvature and the propensity for fusion, too much PE (or too little PS) could result in membranes

B

C

WT Psd1 O

/E

48 kDTom40

s-Mgm1100 kD

75 kDl-Mgm1

63 kDPsd1-3HA

A

Psd1 O/E

WT

84

with too much curvature, either impeding fusion or promoting fission. This finding strongly

implicates the composition of the mitochondrial membrane as a critical regulatory factor in

mitochondrial membrane dynamics. PE is not simply a pro-fusion lipid - the equilibrium

between PE and its precursor, PS, might fine-tune the balance between adequate fusion and

insufficient fusion (or increased fission). It is intriguing that a lipid known for its fusogenic

properties may impede organelle fusion or promote organelle fission. The next phase of

discovery will be to examine how it does so and to determine its mechanism of action.

Continuing with yeast as a model system, one can investigate whether the Psd1 overexpression

phenotype is a fusion defect by employing the same in vivo mitochondrial fusion assay described

in section 3.3.2. To confirm if fusion is impaired, Psd1 can also be overexpressed in a

mitochondrial fission mutant. As discussed in section 1.1.1, decreased mitochondrial fusion can

be rescued by reducing mitochondrial fission; hence, if the overexpression of Psd1 impairs

mitochondrial fusion, its overexpression in a mitochondrial fission mutant should result in

tubular mitochondria similar to that of wild type (WT) cells.

Given that PE might play a role both in mitochondrial fusion and mitochondrial fission, it

would be immensely helpful to have a better knowledge of how PE production is regulated.

Although PE is the second most abundant phospholipid in the cell (after phosphatidylcholine

[PC]), the regulation of its biosynthesis remains largely uncharacterized. Mitochondria interact

with the endoplasmic reticulum (ER) at contact sites known as mitochondrial-associated ER

membranes (MAMs) (291). These points of contact are maintained by the ER-mitochondria

encounter structure (ERMES) (280), and serve critical functions in lipid biosynthesis (292).

MAMs not only act as sites of lipid transfer - they are also enriched in lipid-synthesizing

enzymes (291). The synthesis of PE involves the MAM-enriched enzyme PS synthase (292) that

85

converts cytidyldiphosphate diacylglycerol (CDP-DAG) to PS. PS from the ER must then be

transferred to the outer mitochondrial membrane (OMM) and then to the inner mitochondrial

membrane (IMM) where PS decarboxylase (PSD) resides (293). The transfer of PS from the

OMM to the IMM likely occurs at mitochondrial IMM-OMM contact sites (294,295) maintained

by the recently identified mitochondrial contact site (MICOS) complex (296). Since MAMs and

IMM-OMM contact sites facilitate lipid transfer, they could also regulate lipid biosynthesis. It

was recently reported that the synthesis of PC is regulated by contact sites between the plasma

membrane (PM) and the ER (PM-associated ER [pmaER]) (297). It is proposed that PM-ER

contact sites provide spatial regulation by bringing the lipid substrate and its modifying enzyme

into close proximity (297). Perhaps Psd1/PISD localize to IMM-OMM contact sites in close

proximity to imported PS. This localization would also facilitate the export of PE from

mitochondria. To determine the specific submitochondrial localization of Psd1/PISD,

immunogold electron microscopy (EM) and subcellular fractionation experiments can be

performed. Furthermore, if the MICOS complex is important in PE biosynthesis, disrupting the

MICOS complex will result in alterations in the lipid composition of the mitochondrial

membrane. To analyze the mitochondrial lipid composition of MICOS mutants with disrupted

IMM-OMM contacts, mitochondria can be purified, and lipids can be extracted and analyzed by

thin-layer chromatography and mass spectrometry.

5.2.2 Mitochondrial lipids in mitochondrial biology

The role of lipids in mitochondrial dynamics is a burgeoning area of research. In the last

few years, the role of lipids in mitochondrial membrane fusion has been the intensively studied.

The results from several studies, including my own, indicate that the processing of Mgm1, a key

86

mitochondrial fusion protein, is affected by lipid metabolism. Specifically, the enzymes that

regulate PE and CL levels have been shown to alter s-Mgm1 biogenesis (98-100). Mgm1 exists

as two cleavage isoforms, long-Mgm1 (l-Mgm1) and s-Mgm1. As discussed in sections 1.1.3.2

and 3.3.7, Mgm1 contains two N-terminal hydrophobic regions; l-Mgm1 is formed when the first

hydrophobic region is inserted into the IMM whereas s-Mgm1 is formed when full-length Mgm1

(FL-Mgm1) is further inserted into the membrane by its second hydrophobic region and

subsequently cleaved by the rhomboid intramembrane protease Rbd1/Pcp1 (Figure 1-3). Since

the insertion of the second hydrophobic region of FL-Mgm1 is ATP-dependent, the simple

explanation for the role of PE and CL in regulating s-Mgm1 biogenesis might be through the

regulation of mitochondrial ATP levels, as both PE and CL have been shown to interact with and

influence the activity of the supercomplexes in the electron transport chain (ETC) (104,105,255).

However, since Mgm1 undergoes alternative topogenesis, an exciting possibility is that PE and

CL could reduce the propensity of the first hydrophobic region in Mgm1 to remain in the IMM.

This would increase the likelihood that the second hydrophobic region will be inserted into the

IMM, thereby promoting the formation of s-Mgm1. PE, CL and Mgm1 localize to OMM-IMM

contact sites (294,295,298). It is proposed that prohibitins (PHBs) act as scaffolds to provide

high local concentrations of PE and CL (99,101), without which, mitochondria appear

fragmented (99). This strongly suggests that a specific local amount of PE and CL is required

for normal mitochondrial morphology. Perhaps the high local concentrations of PE and CL that

are achieved by the PHBs maintain mitochondrial morphology by ensuring the right balance of l-

Mgm1 and s-Mgm1. It is well-accepted that alternative topogenesis of Mgm1 is dependent on

ATP, the mitochondrial protein import machinery and the hydrophobicity of the first

hydrophobic region within Mgm1 (81). However, its dependence on the composition of the

87

mitochondrial membrane has not been explored. With recent advances in molecular dynamics

(MD) simulations, this area can now be investigated. However, to draw a strong conclusion

from MD simulations, the crystal structure of l-Mgm1 first needs to be obtained, and might

prove to be the biggest challenge in this area of study. Although l-Mgm1 has been purified (85),

its crystal structure has not been reported. An alternative to obtaining the crystal structure of l-

Mgm1 would be to model its structure on that of a similar dynamin or dynamin-related protein

(DRP) that has been crystallized. However, only the guanosine triphosphate hydrolase (GTPase)

domain of Mgm1 has a high sequence homology to dynamins and DRPs (85). Furthermore, no

crystallized dynamin or DRP has a transmembrane (TM) segment. Hence, it would be difficult

to accurately model the structure of l-Mgm1 on that of other crystallized dynamins or DRPs. l-

Mgm1 lacks a functional GTPase domain in vitro (85) and in vivo (84), and it is proposed that

the membrane insertion of l-Mgm1 restricts its GTPase activity - treating liposome-reconstituted

l-Mgm1 with a detergent resulted in observable l-Mgm1 GTPase activity (85). These data

suggest that the membrane insertion of l-Mgm1 plays an important role in regulating its function.

Thus, determining whether the lipid composition affects l-Mgm1 membrane insertion might shed

light on a possible mechanism of Mgm1 regulation. If true, this will be a novel regulatory

mechanism of mitochondrial membrane fusion.

5.2.3 The specific role(s) of mitochondrial phosphatidylethanolamine in

autophagy

Although a relatively recent discovery, it is well established that mitochondrial dynamics

and function play crucial roles in autophagy (106,108,109). Despite these findings, little is

known about the specific function of the mitochondrial membrane in autophagy. It was reported

88

that mitochondria contribute membranes to growing autophagosomes during starvation-induced

autophagy (111), however, the membrane origin of autophagosomes remains highly debated as

other studies have shown that organelles such as the ER (145,146) and the PM (173) also supply

membranes to growing autophagosomes (121). To begin addressing the role of the

mitochondrial membrane in autophagy, the functional importance of the autophagosomal

membrane first needs to be determined. A previous report demonstrated that the efficiency of

autophagy-related 8 (Atg8) conjugation to PE increases with increasing amounts of PE within

the membrane bilayer, peaking at ~70% PE in an in vitro reconstitution system (189). Strikingly,

mitochondria are the organelles that typically contain the highest amounts of PE in the cell. This

suggests that mitochondria, although not necessarily the only source, could be the preferred

source of membranes for growing autophagosomes due to its higher PE content.

Apart from the amount of PE, the type of PE could also differ between mitochondrial PE

and the PE in other membranes. The majority of mitochondrial PE is synthesized by the

decarboxylation of PS by PSD in the IMM; the Kennedy pathway contributes only a small

amount of PE to mitochondria (229,245,293). Interestingly, mammalian PSD has been shown to

preferentially decarboxylate certain species of PS (246,299), and PSD-derived PE usually

contain polyunsaturated acyl chains, whereas PE derived from the Kennedy pathway usually

contain mono- or di-unsaturated acyl chains (245). The increase in polyunsaturated acyl chains

may alter the distribution and/or packing of PE within the mitochondrial membrane and,

although the biological significance of the different acyl chains is still unknown, one cannot rule

out the possibility that these differences could influence autophagy. In support of this, my

finding that the loss of Psd1 results in a reduction in vacuolar autophagic bodies suggests that the

fusion of autophagosomes with the vacuole/lysosomes could be dependent on certain properties

89

of the autophagosomal membrane. Given that PE is known to play an important role in

membrane fusion and remodelling, it would be consistent with its role as a fusogenic lipid if PE

facilitates the fusion of autophagosomes with the vacuole/lysosomes. To test this hypothesis, a

tandem-tagged mRFP-GFP-LC3 fusion protein can be introduced into mammalian cells

(300,301), and autophagy can be induced by starvation. This assay monitors autophagosome

fusion with lysosomes and takes advantage of lysosomal quenching of GFP but not RFP. Upon

the induction of autophagy, autophagosomes fluoresce yellow due to the co-localization of

mRFP and GFP. However, autophagosomes that have fused with lysosomes will fluoresce red

since the acidic lysosomes quench GFP fluorescence, resulting in only detectable mRFP

fluorescence. If mitochondrial PE is important for the fusion of autophagosomes with

lysosomes, cells with reduced mitochondrial PE should have fewer red autophagosomes

compared to WT cells.

5.2.4 The composition of the mitochondrial membrane in autophagy

The model proposed above is simplistic in that it does not account for other lipids within

the membrane bilayer that could also play important roles in autophagy. Indeed, although PE is

the target lipid for Atg8/LC3, negatively-charged phospholipids also play an important role in

the efficiency of Atg8-PE conjugation (189). The amount of Atg8-PE conjugate formed in the

presence of liposomes containing 20% PE increased when the amounts of negatively-charged

phospholipids, phosphatidylinositol (PI), PS, phosphatidylglycerol (PG) or phosphatidic acid

(PA) were increased (189). This strongly suggests that the composition of the membrane bilayer

plays an important role in autophagy, and that different lipid compositions within the membrane

bilayer may influence Atg8-PE conjugation and perhaps even autophagosome formation.

90

Besides negatively-charged phospholipids, membrane curvature could also play a role in

Atg8-PE conjugation efficiency by influencing the E3-like enzyme, the Atg12-Atg5/Atg16

complex. Atg12-Atg5/Atg16-dependent Atg8-PE conjugation was more efficient in the presence

of giant unilamellar vesicles (GUVs) than small unilamellar vesicles (SUVs) (158). In this in

vitro system, GUVs were less abundant and had less curvature than SUVS, prompting the

authors to propose that membrane availability and/or membrane curvature could affect Atg12-

Atg5/Atg16-dependent Atg8-PE conjugation efficiency (158). Although this experiment was

performed in vitro, the use of GUVs, not SUVs, recapitulated previous in vivo observations that

Atg8-PE conjugation is strictly dependent on the Atg12-Atg5/Atg16 complex (140,158,160),

lending more credibility to this result. The postulation that autophagosomes might have

increased curvature is consistent with the requirement for the fusion of autophagosomes with the

vacuole/lysosomes. Lipids that induce membrane curvature have been shown to promote

membrane fusion (273). Together, these findings indicate that the membrane composition plays

an important role in autophagy, as lipids other than PE can modulate Atg8-PE conjugation

efficiency.

In addition to PI, PS, PG and PA, the mitochondrial membrane also contains CL, a

mitochondrial-specific phospholipid that carries two negative charges and induces increased

membrane curvature due to the size of its hydrophilic head relative to its hydrophobic tail.

Hence, CL fulfills both criteria discussed above: it is negatively-charged and promotes increased

membrane curvature. A well-characterized role of CL is its role in the initiation of apoptosis, a

highly conserved mechanism of programmed cell death (reviewed in reference (302)). Under

normal conditions, CL in the IMM interacts strongly with cytochrome c via electrostatic and

hydrophobic interactions, keeping cytochrome c in the intermembrane space (IMS) tightly

91

associated with the IMM (303-305). Upon apoptotic stimuli, CL is peroxidized and translocates

from the IMM to the OMM (306,307). Although the significance of CL translocation is

unknown, it is proposed that peroxidized CL on the OMM might serve as a signal that indicates

damaged mitochondria (306). The peroxidation of CL facilitates the opening of the

mitochondrial permeability transition pore (MPTP) and the release of cytochrome c (308) from

the mitochondrial IMS to the cytosol to initiate the caspase cascade and consequently, apoptosis

(303).

Interestingly, the translocation of CL from the IMM to the OMM is not unique to

apoptosis. Recently, CL translocation was shown to be important for the recruitment of LC3

during mitophagy, the selective degradation of damaged mitochondria by autophagy (288).

Mutating the residues required for CL binding on LC3 completely abolished mitochondrial co-

localization with autophagosomes, indicating severely impaired mitophagy (288). CL has also

been implicated in general autophagy (287). The immunity-related GTPase, IRGM, translocates

to mitochondria, selectively binds to CL and induces autophagy (287). Although these findings

do not pinpoint the specific role of CL, they strongly suggest that the mitochondrial membrane

might play an important role during autophagy. Although PE, PA and CL have individually

been shown to play important roles in autophagy, I propose that the specific composition of

mitochondrial phospholipids confer favourable properties for Atg8-PE conjugation (or LC3-II

formation) and autophagosome fusion with the vacuole/lysosomes, and hence could be the

preferred source of membranes for growing autophagosomes. As discussed above, CL has

previously been shown to influence autophagy, although its specific role remains unclear. To

determine if CL influences the efficiency of LC3 conjugation to PE, CL levels can be reduced by

knocking down CL synthase, and LC3-I conversion to LC3-II can be monitored by western

92

blotting after the induction of autophagy by starvation. A reduction in the amount of LC3-II in

cells with reduced CL levels would suggest impaired LC3-II formation or increased LC3-II

degradation. To distinguish between the two, this experiment can be repeated in the presence of

Bafilomycin A, a drug that inhibits the fusion of autophagosomes with lysosomes and hence

prevents LC3-II degradation. If mitochondrial CL levels play a role in LC3 conjugation to PE, in

the presence of Bafilomycin A, cells with reduced CL should have reduced levels of LC3-II

compared to WT cells. To determine if autophagosome formation is impaired, GFP-tagged LC3

(GFP-LC3) can be used to monitor autophagosome formation by fluorescence imaging where

autophagosome formation is indicated by GFP-LC3 puncta. If CL influences autophagosome

formation, cells with reduced CL levels should have fewer GFP-LC3 puncta than WT cells.

5.3 Conclusion

In the process of studying the protein components of mitochondrial membrane fusion and

autophagy, several studies have inadvertently discovered important roles of lipids in regulating

these processes. In the last few years, interest in lipid metabolism has resurfaced, and it is

becoming increasingly clear that lipids play crucial roles in modulating protein function and, in

the case of organelle fusion, may influence the kinetics of lipid mixing. The work presented in

this thesis provides new insight into the specific role of mitochondrial-localized PE during

mitochondrial membrane fusion and implicates specific properties of the mitochondrial

membrane during autophagy. Notably, although a reduction in mitochondrial PE compromises

mitochondrial fusion, function and autophagy, it is not essential, as the aforementioned functions

remained intact, albeit with significant defects. Coincidentally, this phenomenon is also

observed for CL - yeast cells lacking CL synthase have reduced mitochondrial activity but still

93

retain mitochondrial function (309) and survive under conditions that necessitate mitochondrial

respiration (310). The overlapping functions of PE and CL may provide a simple explanation for

these observations, as a simultaneous reduction in PE and CL is lethal in yeast (311) and results

in impaired mitochondrial fusion (272). However, these results strongly suggest that a specific

lipid composition is crucial for optimum function, and that the role of mitochondrial lipids in

mitochondrial membrane dynamics and autophagy warrants further study. At the rate that

scientific research is progressing, I have no doubt that the scientific community will continue to

uncover more of the subtle yet significant and important roles that lipids play in the regulation of

mitochondrial membrane dynamics and autophagy; and I look forward to the day that it does.

94

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